This work was supported by the NERC highlights grant (NE/N016777/1). Ensis Ltd, Environmental Scientific Services, Environmental Change Research Centre, University College London sampled and dated the sediment core.

The following SOP provides a step-by-step description of the protocol used to resurrect Daphnia magna dormant eggs, including a detailed description of sampling, isolation of ephippia from the sediment, and establishment of clonal cultures (Figure 1).
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/56637/56637fig1.jpg" /> 
Figure 1: Step-by-step guide to resurrection of Daphnia magna. Sediment from a natural freshwater habitat (A) is sampled with a piston corer (B). The sediment core (C) is sliced in incremental layers of 1 or 0.5 cm (D). Each layer of sediment is stored in a sample zip lock bag (E) in dark and cold conditions (4 &#176;C). Each layer of sediment is weighed and sieved using geological sieves (1 mm and 125 &#181;m mesh sizes, F). White background trays are used to isolate Daphnia magna ephippia (G). Decapsulated dormant eggs (H) are transferred to Petri dishes and exposed to light and temperature stimuli to induce hatching. Hatchlings are transferred to separate jars (I) to establish isoclonal lines. <a href="//cloudfront.jove.com/files/ftp_upload/56637/56637fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
1. Sampling of Sediment Cores
<ol>
	<li>Sample sediment from lakes or ponds using a piston corer. This protocol used Big Ben27, a core tube of approximately 1.5 m in length with an internal tube diameter of 14 cm. Big Ben consists of a piston on a rope and a corer head, to which rods are attached to drive the tube into the sediment. A core catcher aids the support of the core tube when full of sediment. To extrude the sediment, a framework keeps the core tube upright and stationary, and a modified bottle jack is used to push the piston upwards during the extrusion process (Supplementary Video 1).
	<ol>
		<li>For shallow ponds of less than 1 m in depth, use a plexiglass gravity corer of no more than 6 cm of diameter manually pushed into the sediment.</li>
		<li>For deep lakes (&#62;6 m of depth), use Livingston piston corers28 or single-drive Griffith sediment corers with the aids of an anchored pontoon boat. The Livingstone-type drive rod piston corer can be used in water up to about 30 m deep to collect successive one-meter drives of soft to consolidated lake sediment. The single-drive Griffith corer consists of a simple but robust core head that connects standard polycarbonate tubes to Livingstone drive rods. The corers are pushed into the sediment with the rods, and a piston provides the suction needed for the recovery of sediment (Figure 2).</li>
		<li>For retrieving continuous, undisturbed cores, use vibracoring. These corers work on a variety of water depths and can retrieve core samples of different lengths, depending on sediment lithology. Low amplitude vibration that is transferred from the vibracore head down through the attached barrel or core tube liquefies sediments, enabling the core barrel attached to the vibracore unit to penetrate into the liquefied sediments. Some vibracorers are small, lightweight, and portable, others are large heavy units that can only be deployed from large vessels. The choice of corers depends on the lithology of sediment.</li>
	</ol>
	</li>
	<li>Slice the core horizontally in incremental layers of 1 cm or less using a flat metal surface (Supplementary Video 1). Sediment corers like the one used here are designed to reduce hydrostatic pressures at extrusion, reducing disturbance of the sediment layers. When other corers are used, the outer rind of each sediment layer may be removed with a cookie-cutter sort of blade to limit contamination among layers.</li>
	<li>Collect each sediment layer in a separate sampling bag (Supplementary Video 1), and store in dark and cold (4 &#176;C) conditions.</li>
	<li>Collect a minimum of 5 g of sediment from all layers for radiometric dating. As radiometric dating is an established protocol for a detailed description of the dating assay, refer to existing publications12,13.</li>
</ol>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/56637/56637fig2.jpg" /> 
Figure 2: Cartoon of the piston coring procedure. Piston corer, a hollow tube with an internal sliding seal (the piston) that produces a weak vacuum. When the piston touches the sediment-water interface, the weight pushes the core barrel into the sediment and the vacuum causes the sediment being cored to enter and move up the tube without disturbing the sediment layers. <a href="//cloudfront.jove.com/files/ftp_upload/56637/56637fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
2. Sieving of Sediment Layers
<ol>
	<li>Using a precision scale, weigh each sediment layer for future reference. Use the surface area and weight to calculate the species density in the lake.</li>
	<li>Sieve each sediment layer using two geological sieves piled on top of each other. The first sieve has a mesh size of 1 mm and separates clay, large invertebrates and particulate matter, e.g. seeds, plants, and insects, from the remainder of the sediment. The second sieve has a mesh of 125 &#181;m and separates D. manga ephippia and small particulate from the remainder of the sediment (Supplementary Video 2).</li>
	<li>Transfer small aliquots of the sediment fraction collected onto the 125 &#181;m mesh sieve to a white background tray. Depending on the type of sediment, smaller or larger aliquots of sediment may be transferred at each time.
	<ol>
		<li>Add small volumes (up to 200 mL) of medium to the sampling white tray to re-suspend the transferred sediment fraction and facilitate eye spotting of ephippia (Figure 3). The medium used to resuspend the sediment can be dechlorinated tap water, borehole water, COMBO29, or ADaM medium (Aachener Daphnien)30. Hereafter, the term &#39;medium&#39; will be used to refer to either or all the listed solutions.</li>
	</ol>
	</li>
</ol>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/56637/56637fig3.jpg" /> 
Figure 3: Daphnia magna ephippium. Dormant Daphnia magna eggs immediately after decapsulation. The ephippium (A), the inner egg membrane (B), and the dormant eggs (C) are shown. Scale bar = 500 &#181;m.
3. Decapsulation of Ephippia and Hatching
<ol>
	<li>With a disposable Pasteur pipette or using microdissection forceps, transfer individual ephippia to Petri dishes filled with 10 mL of medium. Use at least a Petri dish per layer of sediment.</li>
	<li>Under a stereo microscope, decapsulate each ephippia using microdissection forceps by forcing open the chitin case (Supplementary Video 3). Remove the resting egg inner membrane delicately, paying attention to not disrupt the eggs, and transfer them to the Petri dish filled with medium using a Pasteur pipette. Decapsulation increases hatching success in D. magna; however, it may not be required or it may be challenging for other Daphnia species producing smaller ephippia.</li>
	<li>Expose the decapsulated eggs to a full spectrum long day photoperiod light (16:8 light:dark) and high temperature (20 &#177; 1 &#176;C) to induce hatching in a controlled temperature device (incubator) or room. Hatching occurs between 48 h and several weeks (up to four; Supplementary Video 4). In absence of decapsulation, directly expose the ephippia to hatching stimuli (long day photoperiod light (16:8 light:dark) and high temperature (20 &#177; 1 &#176;C)).</li>
</ol>
4. Establishing Isoclonal Lines of Daphnia magna
<ol>
	<li>Establish isoclonal lines from single hatchlings by transferring individual D. magna from step 3.3 to separate jars filled with 200 mL of medium using a disposable Pasteur pipette. Each individual is genetically distinct, being the result of sexual recombination.</li>
	<li>Maintain isoclonal lines indefinitely in stock conditions consisting of 10 &#177; 1 &#176;C, 16:8 light:dark regime, fed weekly with 0.2 mg C/L of Chlorella vulgaris or other green algae (e.g., Scenedesmus obliquus). Renew the medium every third week. Stock conditions may change with temperature, feeding regimes, and species.</li>
</ol>
5. Key Studies
NOTE: A description of two key studies is provided in which resurrected D.magna (sub)populations from the sedimentary archive of Lake Ring (Denmark) are used to assess the evolutionary response to warming. Three (sub)populations were resurrected from the following time periods: 1960-1970, 1970-1985, and &#62;1999. D. magna hatching success from the sedimentary archive ranged between 11 and 58% (Figure 4). From the hatchlings obtained from each timeperiod, a random subset was chosen for the two key studies described here. These studies were designed to assess whether (sub)populations resurrected from different time periods along the temperature gradient showed differences in fitness-linked life history traits (5.1), and whether they had different competitive abilities (5.2) after exposure to warming.
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/56637/56637fig4.jpg" /> 
Figure 4: Hatching success in a sedimentary archive sampled from Lake Ring. The proportion of successful hatchlings along the sedimentary archive of Lake Ring used in the key studies. <a href="//cloudfront.jove.com/files/ftp_upload/56637/56637fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<ol>
	<li>Common garden experiments
	<ol>
		<li>Transfer ten hatchlings per (sub)population from stock cultures to common garden conditions: 16 &#176;C; long photoperiod (16:8 light:dark regime); feed daily with 0.8 mg C/L of Chlorella vulgaris, and renew medium every second day. Maintain the hatchlings in common garden conditions for at least two generations (ca. 45 days). Common garden conditions reduce interference from the maternal effect and synchronize reproduction among hatchlings.</li>
		<li>Upon release of the second brood into the brood chamber, transfer adult females from the second generation to 500 mL jars filled with medium until they release the second brood of juveniles.</li>
		<li>Randomly transfer individual juveniles of 24&#8211;48 h, born from the second brood of the second generation, to 100 mL jars filled with medium and exposed to the following experimental conditions: 18 &#176;C (current temperature in the lake) and 24 &#176;C (warming, temperature forecasted by the IPCC 26 for the upcoming 100 years), 16:8 light:dark regime, and feed daily 0.8 mg C/L of Chlorellavulgaris.</li>
		<li>On each experimental animal, measure the size at maturity with a stereomicroscope as the distance between the head and the base of the tail spine (Figure 5). Photograph each animal and subsequently analyze its size using a suitable image software.</li>
		<li>Measure the age at maturity: the day in which eggs are observed for the first time in the brood chamber.</li>
		<li>Measure the mortality: the number of extinct individuals during the experiment.</li>
		<li>Measure the fecundity: the total number of offspring released in first and second clonal reproduction.</li>
		<li>Measure the population growth rate, estimated using the Euler&#39;s equation (1): 
		<img alt="Equation" src="/files/ftp_upload/56637/56637eq1.jpg" /> (1) 
		Where lx is the proportion of survival at age x, bx is the number of neonates produced per surviving individual at age x, and r is the intrinsic rate of natural increase.</li>
		<li>Perform statistical analysis using commercially available software. Here, use R31 to plot reaction norms (phenotypic expression of a single genotype across environments) for each life history trait, using the ggplot2 package. Calculate mortality rates per population via survival analysis (R package rms; https://cran.r-project.org/web/packages/rms/rms.pdf). Finally, perform an analysis of variance (ANOVA, Table 2) in R31 to assess whether the effect of temperature on traits can be explained by evolution (differences among populations), plasticity (response to treatment), or evolution of plasticity (population x treatment).</li>
	</ol>
	</li>
	<li>Competition experiment
	<ol>
		<li>Transfer ten hatchlings per population from stock cultures to common garden conditions: 20 &#176;C, long photoperiod (16:8 light:dark regime), feed daily with 0.8 mg C/L of Chlorella vulgaris, and renew medium every second day. Maintain common garden conditions for at least two generations (ca. 45 days) to reduce interference from maternal effects.</li>
		<li>Randomly assign five juveniles of 24 to 48 h from the second brood of the second generations of each hatchling to experimental mesocosms (20 L plastic aquaria filled with medium), in triplicates, at a density of 10 animals/L.</li>
		<li>Expose the mesocosms to 24 &#176;C, 16:8 L:D regime, and feed daily with 0.8 mg C/L of Chlorellavulgaris in a controlled temperature chamber or incubator for a minimum of four weeks (&#62;3 clonal generations).</li>
		<li>Cull each mesocosm weekly, refreshing 10% of the medium to simulate a population dynamic that Daphnia may encounter in the natural environment. Impose culling by removing a known volume of medium and animals from each aquarium (1.2 L in this case), and by replacing the culled volume with fresh medium. Start the culling regime after the experimental animals reach maturity (day 10).</li>
		<li>At the end of the fourth week, sample 32 animals from each mesocosm to assess shifts in genotypic composition as compared to the initial inoculum.
		<ol>
			<li>Place individual Daphnia in microcentrifuge tubes and remove excess liquid with the help of a Pasteur pipette.</li>
			<li>Flash freeze individual tubes in liquid nitrogen and store at -80 &#176;C.</li>
			<li>Extract genomic DNA from single individuals using available protocols and following the manufacturer&#39;s instructions.</li>
			<li>Amplify the extracted DNA using a number of genetic markers sufficient to provide a unique multilocus genotype per hatchling. Here, 8 microsatellites arranged in a single multiplex (M01, Table 1) were used following established protocols32,33.</li>
			<li>Genotype amplified fragments on a fragment analyzer.</li>
			<li>Conduct fragment analysis with a commercial or freely available software using a suitable size standard.</li>
			<li>Assess genotypic composition at the end of the experiment by genotyping the 32 individuals with a set of microsatellite loci as described33, and calculating the frequency of each genotype at the end of the experiment as compared to the initial inoculum.</li>
		</ol>
		</li>
	</ol>
	</li>
</ol>
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/56637/56637fig5.jpg" /> 
Figure 5: Adult female Daphnia magna. Adult female Daphnia magna with parthenogenetic eggs in the brood chamber. The distance between the head and the base of the tail spine is used to measure size of the animal. The red lines indicate the size measurements. Scale bar = 500 &#181;m.

The authors have nothing to disclose

Due to the high thermal conductivity of water, freshwater ecosystems are at higher risk of biodiversity loss than terrestrial ecosystems in the face of global warming34. It is, therefore, critical to understand the response of keystone species in these ecosystems and identify coping mechanisms to survive thermal stress. The understanding of these mechanisms at the species and community levels can help predict how species are impacted by global warming and how the effect on individual species cascades to other trophic levels. Ultimately, understanding mechanisms of responses to global warming enables the identification of remediation strategies to mitigate extinctions.
The case studies presented here show that the response of D. magna to temperature increase is pervasively mediated by plasticity in life history traits and that response to temperature increase alone does not impose clear fitness costs, at least in the population studied here. High plasticity in life history traits is supported by non-significant differences in competing abilities of the (sub)populations in the presence of warming. However, longer-term competition experiments on multiple populations may be necessary to generalize these findings.
Resurrection of dormant stages provides an unprecedented resource to study mechanisms of adaptation and trajectories of a species&#39; evolution through time10. Zooplankton species benefit from a rapid generation time (about 2 weeks), and the viability of dormant stages, which allows an ancestor to compete headtohead against its own descendants, or to &#39;replay&#39; evolution that starts from various past states. Resurrection ecology essentially enables the investigation of whether a particular evolutionary outcome is contingent on some prior event. The identification of the genetic elements of evolution is currently possible in laboratory experiments using microorganisms for which &#39;ancestral lines&#39; are frozen and resuscitated for comparative analysis with their evolved descendant6. However, one of the main limitations of working with laboratory organisms is that the &#39;ancestral state&#39; is an already shifted baseline. The study of dormant stages allows the sampling of specimens from time predating any stress event (e.g., pristine environmental conditions) and to measure evolutionary trajectories from undisturbed environmental conditions to various past states until modern times. In recent years, the study of DNA polymorphism in resurrected or still dormant zooplankton stages has provided important insights into past demographic and adaptive processes that have contributed to the genetic makeup of present-day populations14,16,25,33,35,36. With the higher accessibility of high throughput sequencing technologies, the genome and transcriptome of resurrected or still dormant stages can be sequenced and the type and number of genetic changes accumulated in evolving populations over time measured.
The resurrection SOP presented here has important applications in the field of multi-omics on two levels. Multi-omics technologies can be applied to resurrected specimens, allowing an exhaustive analysis of the molecular elements involved in adaptive responses to environmental selection pressure. In addition, omics technologies can be applied to decapsulated but still dormant stages. So far, the application of high throughput sequencing technologies to resting stages has been limited by the requirement of a large amount of input material. These limitations are being lifted37. With the lowering requirements for input material and progress in nanofluidics, whole genome sequencing (WGS) is now possible from as little as 1 ng or a few pg of starting material38. The use of whole genome amplification (WGA) and whole transcriptome amplification (WTA) techniques, enabling the enrichment of DNA and RNA from very small amounts of tissue, has revolutionized both metagenomics39,40 and medical research41. These technologies applied to decapsulated dormant eggs enable the exceeding of limitations associated with viability of dormant stages and the investigating of extended time periods (e.g., centuries).
The resurrection of invertebrate communities producing resting stages enables the alignment of community histories with known changes in the natural landscapes, or with environmental changes inferred from analyses of the sediments orsoils2. The analysis of community changes in response to environmental change provides us with the ability to quantify eco-evolutionary feedbacks42 that have substantial consequences on population persistence43, trophic interactions44, community assembly45, and changes in ecosystem functions and services46. Finally, accurate predictions about biological responses to environmental change are paramount to guide the protection of biodiversity47. Current predictive models are inaccurate in this respect because they do not take into account important biological mechanisms such as demography, dispersal, evolution, and species interactions. Understanding how these processes change over time and using this information as a prior in forecast modelling will improve our ability to predict species and community persistence in the face of environmental change2.
The application of the SOP presented here is not without challenges. The primary limitation of resurrecting dormant stages is the need of specialized equipment for sampling. Additionally, the entire process, from sediment sieving to establishment of clonal cultures, requires considerable hands-on time.
Some of the SOP steps presented here are readily transferable to other Daphnia species. These are: sampling, establishment of clonal lines, and experimental design. However, other steps of the SOP may require further optimization tailored to the species under study. Decapsulation is often applied to D. magna specimens to improve hatching success. However, this approach may not be suitable for smaller specimens. Hatching stimuli can also vary among species48 and among conspecific specimen49. Hence, an ad hoc optimization of the hatching steps of the SOP may be required prior to applications to other crustaceans. Whilst the hatching success of the D. magna population resurrected from Lake Ring (30.5% across the sedimentary archive) is in line with previous results49, hatching success varies with the preservation state of the sediment, the species50,51, and the geographic origin of the sediment48. Future studies on the mechanisms that regulate entry and progression through the phases of diapause is required to identify optimal hatching stimuli tailored to different species.
Finally, background knowledge of the study system, in particular the presence of the species of interest over time, is advisable. This can be achieved via historical records. If historical records are not available, sampling and screening of the surface layers of the lake sediment prior to core sampling is advisable, although it may provide information only on the most recent history.

Long-term studies are critical to understanding ecological and evolutionary processes in nature and in assessing how species respond to and persist during environmental change1. This is because eco-evolutionary processes happen across generations and changes in the environment occur over long time spans. Furthermore, long-term studies provide key empirical data that improve the accuracy of predictive modeling to forecast evolutionary responses of natural ecosystems to environmental changes2. The accuracy of these models is critical to implement management and conservation strategies to preserve biodiversity and ecosystem services.
Excluding a few exceptional cases (e.g., Galapagos Darwin finches3 and algae4), long-term studies are largely limited to species with short generation time that can be propagated in the laboratory5,6,7,8. Hence, the processes underpinning evolutionary dynamics remain elusive. Because of logistic difficulties associated with accessing temporal samples, empirical data are studied more frequently in a spatial than in a temporal context, and temporal eco-evolutionary processes are inferred or modeled from spatial data. This approach is known as &#39;space-for-time&#39; substitution9, whereby space is adopted as a surrogate to study temporal evolutionary dynamics. The main limitation of the &#39;space-for-time&#39; substitution is that rates of adaptation at different spatial scales differ from temporal variation in the same population; hence, inferences based on replacement of time with space are biased10.
A powerful alternative that allows studying evolutionary dynamics in natural ecosystems over time is the analysis of ecological and genetic changes in species producing dormant stages11. These dormant stages accumulate to form stratified biological archives that can be accurately dated and paleolimnologically characterized12,13. Importantly, these dormant stages can be resuscitated and used in laboratory experiments, where their evolutionary response to environmental change can be directly measured. Historical populations can be competed against their modern evolved descendants to study fitness changes and the function of genes evolving in step with environmental change14,15,16.
Dormant stages include seeds, cysts, spores, and egg banks. Although the first studies on resuscitated dormant eggs dates back to the late 1980s17, and a handful of studies have applied this technique in the early 1990s18,19, the field of Resurrection Ecology has been formally established by the seminal paper of Kerfoot and co-workers in 199920. This practice has been applied mainly in paleolimnological reconstructions of freshwater species17,21,22. However, a SOP is not yet available. Here, a step-by-step description of the resurrection protocol applied to dormant eggs of the zooplankton species Daphnia magna is provided, from the sampling of sediment to the establishment of clonal cultures from hatchlings. Steps of the SOP that are readily transferable to other species of Daphnia, as well as steps that may require additional optimization, are discussed.
Daphnia are freshwater zooplankters present in the majority of lotic habitats23 . Daphnia species are either obligate asexual or cyclical parthenogens. D. magna is a cyclical parthenogen that reproduces clonally under favorable environmental conditions24. When environmental conditions deteriorate, male production occurs and sexual recombination leads to the formation of fertilized eggs that enter a state of dormancy protected from the environment by a chitin case called ephippium. A proportion of these dormant eggs hatch when favorable environmental conditions return. However, a large proportion of the dormant egg bank never has a chance to hatch and thus build up biological archives over time. Dormant stages remain buried in the sediment of lakes and ponds and can be resurrected for the study of evolutionary dynamics over extended time periods. Because dormant eggs of D. magna are the result of sexual recombination, they are a good representation of the natural genetic diversity of the species25. Moreover, they can be maintained via clonal reproduction in the laboratory. These characteristics provide the unique advantage of isogenic model organisms, while retaining the natural genetic diversity.
Two key studies are presented to demonstrate the advantages of directly comparing historical and modern descendants of the same population of D. magna experiencing environmental selection pressure over time. D. magna specimens were resurrected from Lake Ring (Denmark), a shallow (5 m depth; surface 22 ha) mixed pond that has experienced an increase in average temperature and heat waves occurrence over time. D. magna (sub)populations were resurrected along this temporal gradient spanning 60 years (1960&#8211;2005) and studied to investigate evolutionary response to temperature warming. In the first study in a common garden experiment, changes in fitness-linked life history traits were measured in response to an increase in temperature of +6 &#176;C, in line with the predictions of the Intergovernmental Panel for Climate Change for the upcoming 100 years26. In the second study, a mesocosm experiment was used to measure the competitive abilities of the three (sub)populations under warming. These experiments combined show that in presence of warming as the only stress, all life history traits and populations show a high level of plasticity and have equal competitive abilities. These findings suggest that warming as a single stress does not impose significant fitness costs, at least in the population studied here.

<table><tbody><tr><td>sampling bags</td><td>Fisher Scientific</td><td>11542783</td><td>Sampling bag revolve round wires closure system and safety tabs sterile polyethylene with writing area clear 89&amp;micro;m thickness 140mm x 229mm, 720mL Fisherbrand</td></tr><tr><td>piston corer</td><td>ENSIS ltd</td><td>na</td><td>Long, heavy tube plunged into the sea, lake, pond floor to extract samples of mud sediment. Piston corers have a viariable diameter and are generally in PVC</td></tr><tr><td>precision scale</td><td>Veritas-M124A</td><td>TLP-50</td><td>Analytical Balance</td></tr><tr><td>geological sieve</td><td>UKGE limited</td><td>SV7521</td><td>200 mm diameter geological sieve - 1 mm mesh</td></tr><tr><td>geological sieve</td><td>UKGE limited</td><td>SV7525</td><td>200 mm diameter geological sieve - 0.125 mm mesh</td></tr><tr><td>white sampling tray</td><td>nhbs</td><td>http://www.nhbs.com/title/view/159614?ad_id=1509</td><td>Standard mulipurpose lab trays</td></tr><tr><td>pasteur pipette</td><td>Globe Scientific inc.</td><td>138020B</td><td>Transfer Pipet, 1.7mL, General Purpose, 87mm, Bulb Draw - 0.9mL</td></tr><tr><td>stereo microscope</td><td>nikon</td><td>smz800</td><td>Microscope with magnification range 1x -8x linked to a camera control unit</td></tr><tr><td>petri dish</td><td>EduLab</td><td>153-533</td><td>Sterile 90mm diameter plastic petri dish</td></tr><tr><td>glass jars</td><td>compak</td><td>Round Jam Jars 4oz</td><td>100 mL jars</td></tr><tr><td>glass jars</td><td>compak</td><td>Atum Jars/ Bonta Jar 10oz</td><td>200 mL jars</td></tr><tr><td>glass jars</td><td>bottlecompanysouth</td><td>500ml Food Jam Jar With Twist Off Lid</td><td>500 mL jars</td></tr><tr><td>statistical software R</td><td>https://cran.r-project.org/</td><td>na</td><td>Free online GNU&amp;nbsp; language and environment for statistical computing and graphics</td></tr><tr><td>microdissection forceps</td><td>Fisher Scientific</td><td>41122405</td><td>Fine point stainless steel forceps for microdissections</td></tr><tr><td>image software</td><td>https://imagej.nih.gov/ij/index.html</td><td>na</td><td>Open source ImageJ image processing toolkit written in Java</td></tr><tr><td>mesocosm</td><td>amazon</td><td>na</td><td>Nobby Fauna-Box III, 41 x 23 x 29 cm, 20.0 Liter</td></tr><tr><td>mirocentrifuge tubes</td><td>Sigma_Aldrick - Merck</td><td>Z606340</td><td>premium microcentrifuge tubes 1.5 mL</td></tr><tr><td>AGENCOURT DNAdvance</td><td>Beckman Coulter</td><td>A48705</td><td>DNA extraction kit</td></tr><tr><td>size standard</td><td>Thermo Fisher Scientific</td><td>4322682</td><td>LIZ500 - Size standard compatible with ABI sequencers</td></tr><tr><td>ABI3032 sequencer</td><td>ABI</td><td>na</td><td>Sequencer used to perform fragment analysis or sanger sequencing</td></tr></tbody></table>

resurrection of dormant daphnia magna: protocol and applications

Long-term studies enable the identification of eco-evolutionary processes that occur over extended time periods. In addition, they provide key empirical data that may be used in predictive modelling to forecast evolutionary responses of natural ecosystems to future environmental changes. However, excluding a few exceptional cases, long-term studies are scarce because of logistic difficulties associated with accessing temporal samples. Temporal dynamics are frequently studied in the laboratory or in controlled mesocosm experiments with exceptional studies that reconstruct the evolution of natural populations in the wild.
Here, a standard operating procedure (SOP) is provided to revive or resurrect dormant Daphnia magna, a widespread zooplankton keystone species in aquatic ecosystems, to dramatically advance the state-of-the-art longitudinal data collection in natural systems. The field of Resurrection Ecology was defined in 1999 by Kerfoot and co-workers, even though the first attempts at hatching diapausing zooplankton eggs date back to the late 1980s. Since Kerfoot&#39;s seminal paper, the methodology of resurrecting zooplankton species has been increasingly frequently applied, though propagated among laboratories only via direct knowledge transfer. Here, an SOP is described that provides a step-by-step protocol on the practice of resurrecting dormant Daphnia magna eggs.
Two key studies are provided in which the fitness response of resurrected Daphnia magna populations to warming is measured, capitalizing on the ability to study historical and modern populations in the same settings. Finally, the application of next generation sequencing technologies to revived or still dormant stages is discussed. These technologies provide unprecedented power in dissecting the processes and mechanisms of evolution if applied to populations that have experienced changes in selection pressure over time.

Long-term empirical data are critical to the understanding of evolutionary dynamics and persistence of natural populations. Such data are generally challenging to obtain because of logistic difficulties associated with accessing temporal samples and the requirement of committing long-term to data collection. In the two key studies presented here, empirical evidence of the response to temperature of a central zooplankter in freshwater ecosystems is provided over evolutionary times. This is enabled by the use of layered dormant egg banks that provide the opportunity to study the response of historical populations and their modern descendants to environmental stress in common experimental settings.
Common garden experiment 
The common garden experiment showed that all life history traits responded to temperature (Figure 6 and Figure 7). The ANOVA analysis revealed that all (sub)populations respond to temperature via plasticity (Table 2), except for mortality, which is unresponsive. Evidence of evolutionary changes (differences among (sub)populations) was observed only in population growth rate (Table 2), which significantly increased in two of the three (sub)populations at 24 &#176;C (Figure 6).
<img alt="Figure 6" class="xfigimg" src="/files/ftp_upload/56637/56637fig6.jpg" /> 
Figure 6: Common garden experiment. Reaction norms for life history traits (fecundity, size, and age at maturity) and population growth rate (r) are shown for each (sub)population under temperature warming (24 &#176;C) as compared to the common garden and current temperature regime (18 &#176;C). The population growth rate &#39;r&#39; is calculated using the Eulerian equation (1). Confidence intervals are shown. (Sub)populations are color coded: (i) blue: 1960-1970; (ii) green: 1970&#8211;1985; (iii) red: &#62;1999. <a href="https://cloudflare.jove.com/files/ftp_upload/56637/56637fig6large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 7" class="xfigimg" src="/files/ftp_upload/56637/56637fig7.jpg" /> 
Figure 7: Mortality. Mortality rates per (sub)population (1960-1970; 1970-1985; &#62;1999) are shown under warming (24 &#176;C) as compared to modern temperature regimes (18 &#176;C). <a href="https://cloudflare.jove.com/files/ftp_upload/56637/56637fig7large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Mesocosm experiment 
After four weeks of selection, represented by warming at 24 &#176;C, the frequency of the three (sub)populations did not change significantly (&#967;2 = 0.55, P = 0.76) as compared to the initial inoculum (Figure 8). Among the 30 genotypes inoculated in the mesocosm experiment, the majority was identified after four weeks of selection (Figure 9). Specifically, 70% of the inoculated genotypes were recovered, compatible with Poissonian expectation of recovering at least one representative of each genotype in a sample of 32 individuals.
<img alt="Figure 8" class="xfigimg" src="/files/ftp_upload/56637/56637fig8.jpg" /> 
Figure 8: Competition experiment - population frequency. Population-averaged median and quartiles (25th and 75th), is shown for the three (sub)populations of D. magna after four weeks of selection in mesocosm competition experiments (24 &#176;C), as compared to an initial equal frequency (at the start). (Sub)populations are color coded as shown in Figure 6. <a href="https://cloudflare.jove.com/files/ftp_upload/56637/56637fig8large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 9" class="xfigimg" src="/files/ftp_upload/56637/56637fig9.jpg" /> 
Figure 9: Competition experiment - genotype frequency. Genotype frequencies &#8212; averaged median and quartiles (25th and 75th), are shown after four weeks of exposure to warming (24 &#176;C) as compared to an initial equal frequency of genotypes (dotted line). Names on the x-axis are the inoculated genotypes ID, grouped per (sub)population (blue, 1960-1970; green, 1970-1985; red, &#62;1999). <a href="https://cloudflare.jove.com/files/ftp_upload/56637/56637fig9large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Locus</td>
			<td>AN</td>
			<td>Size range (bp)</td>
			<td colspan="2">Primers (5&#8217;-3&#8217;)</td>
			<td>Dye label</td>
			<td>Repeat motif</td>
			<td>Tm</td>
		</tr>
		<tr>
			<td rowspan="2">B008</td>
			<td rowspan="2">HQ234154</td>
			<td rowspan="2">150&#8211;170</td>
			<td colspan="2">F: TGGGATCACAACGTTACACAA</td>
			<td rowspan="2">VIC</td>
			<td rowspan="2">(TC)9</td>
			<td rowspan="2">56</td>
		</tr>
		<tr>
			<td colspan="2">R: GCTGCTCGAGTCCTGAAATC</td>
		</tr>
		<tr>
			<td rowspan="2">B030</td>
			<td rowspan="2">HQ234160</td>
			<td rowspan="2">154&#8211;172</td>
			<td colspan="2">F: CCAGCACACAAAGACGAA</td>
			<td rowspan="2">PET</td>
			<td rowspan="2">(GA)11</td>
			<td rowspan="2">56</td>
		</tr>
		<tr>
			<td colspan="2">R: ACCATTTCTCTCCCCCAACT</td>
		</tr>
		<tr>
			<td rowspan="2">B045</td>
			<td rowspan="2">HQ234168</td>
			<td rowspan="2">118&#8211;126</td>
			<td colspan="2">F: GCTCATCATCCCTCTGCTTC</td>
			<td rowspan="2">NED</td>
			<td rowspan="2">(TG)8</td>
			<td rowspan="2">56</td>
		</tr>
		<tr>
			<td colspan="2">R: ATAGTTTCAGCAACGCGTCA</td>
		</tr>
		<tr>
			<td rowspan="2">B050</td>
			<td rowspan="2">HQ234170</td>
			<td rowspan="2">234&#8211;248</td>
			<td colspan="2">F: TTTCAAAAATCGCTCCCATC</td>
			<td rowspan="2">6FAM</td>
			<td rowspan="2">(GAA)6</td>
			<td rowspan="2">56</td>
		</tr>
		<tr>
			<td colspan="2">R: TATGGCGTGGAATGTTTCAG</td>
		</tr>
		<tr>
			<td rowspan="2">B064</td>
			<td rowspan="2">HQ234172</td>
			<td rowspan="2">135&#8211;151</td>
			<td colspan="2">F: CTCCTTAGCAACCGAATCCA</td>
			<td rowspan="2">6FAM</td>
			<td rowspan="2">(TC)8</td>
			<td rowspan="2">56</td>
		</tr>
		<tr>
			<td colspan="2">R: CAAACGCGTTCGATTAAAGA</td>
		</tr>
		<tr>
			<td rowspan="2">B074</td>
			<td rowspan="2">HQ234174</td>
			<td rowspan="2">196&#8211;204</td>
			<td colspan="2">F: TCTTTCAGCGCACAATGAAT</td>
			<td rowspan="2">NED</td>
			<td rowspan="2">(GT)9</td>
			<td rowspan="2">56</td>
		</tr>
		<tr>
			<td colspan="2">R: TGTGTTCCTTGTCAACTGTCG</td>
		</tr>
		<tr>
			<td rowspan="2">B096</td>
			<td rowspan="2">HQ234181</td>
			<td rowspan="2">234&#8211;240</td>
			<td colspan="2">F: GGATCTGGCAGGAAGTGGTA</td>
			<td rowspan="2">VIC</td>
			<td rowspan="2">(AC)15</td>
			<td rowspan="2">56</td>
		</tr>
		<tr>
			<td colspan="2">R: TTGAACCACGTCGAGGATTT</td>
		</tr>
		<tr>
			<td rowspan="2">B107</td>
			<td rowspan="2">HQ234184</td>
			<td rowspan="2">250&#8211;274</td>
			<td colspan="2">F: GGGGTGAAGCATCAAAGAAA</td>
			<td rowspan="2">PET</td>
			<td rowspan="2">(CT)8</td>
			<td rowspan="2">56</td>
		</tr>
		<tr>
			<td colspan="2">R: TGTGACCAGGATAAGAGAAGAGG</td>
		</tr>
	</tbody>
</table>
Table 1: Microsatellite multiplex. The NCBI Accession Number (AN), the multiplex information, the PCR primer sequences, the PCR size range, the repeat motif, the dye used to label the forward primer, and the annealing temperature (Tm) are shown.
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td colspan="2">Pop Growth rate (r)</td>
			<td>Df</td>
			<td>F</td>
			<td>P</td>
		</tr>
		<tr>
			<td colspan="2">Evolution (Pop)</td>
			<td>2</td>
			<td>30.309</td>
			<td>&#60;0.001</td>
		</tr>
		<tr>
			<td colspan="2">Plasticity (Temp)</td>
			<td>1</td>
			<td>531.546</td>
			<td>&#60;0.001</td>
		</tr>
		<tr>
			<td colspan="2">Evol. Plasticity (Pop x Temp)</td>
			<td>2</td>
			<td>65.137</td>
			<td>&#60;0.001</td>
		</tr>
		<tr>
			<td colspan="2">Mortality</td>
			<td>Df</td>
			<td>F</td>
			<td>P</td>
		</tr>
		<tr>
			<td colspan="2">Evolution (Pop)</td>
			<td>2</td>
			<td>2.234</td>
			<td>0.1162</td>
		</tr>
		<tr>
			<td colspan="2">Plasticity (Temp)</td>
			<td>1</td>
			<td>2.679</td>
			<td>0.1071</td>
		</tr>
		<tr>
			<td colspan="2">Evol. Plasticity (Pop x Temp)</td>
			<td>2</td>
			<td>1.8657</td>
			<td>0.164</td>
		</tr>
		<tr>
			<td colspan="2">Fecundity&#160;</td>
			<td>Df</td>
			<td>F</td>
			<td>P</td>
		</tr>
		<tr>
			<td colspan="2">Evolution (Pop)</td>
			<td>2</td>
			<td>1.8852</td>
			<td>0.1633</td>
		</tr>
		<tr>
			<td colspan="2">Plasticity (Temp)</td>
			<td>1</td>
			<td>6.8934</td>
			<td>0.0117</td>
		</tr>
		<tr>
			<td colspan="2">Evol. Plasticity (Pop x Temp)</td>
			<td>2</td>
			<td>1.6511</td>
			<td>0.203</td>
		</tr>
		<tr>
			<td colspan="2">Size at maturity</td>
			<td>Df</td>
			<td>F</td>
			<td>P</td>
		</tr>
		<tr>
			<td colspan="2">Evolution (Pop)</td>
			<td>2</td>
			<td>0.211</td>
			<td>0.8106</td>
		</tr>
		<tr>
			<td colspan="2">Plasticity (Temp)</td>
			<td>1</td>
			<td>11.1361</td>
			<td>0.0017</td>
		</tr>
		<tr>
			<td colspan="2">Evol. Plasticity (Pop x Temp)</td>
			<td>2</td>
			<td>0.6586</td>
			<td>0.5225</td>
		</tr>
		<tr>
			<td colspan="2">Age at maturity</td>
			<td>Df</td>
			<td>F</td>
			<td>P</td>
		</tr>
		<tr>
			<td colspan="2">Evolution (Pop)</td>
			<td>2</td>
			<td>0.7811</td>
			<td>0.4637</td>
		</tr>
		<tr>
			<td colspan="2">Plasticity (Temp)</td>
			<td>1</td>
			<td>8.0764</td>
			<td>0.0066</td>
		</tr>
		<tr>
			<td colspan="2">Evol. Plasticity (Pop x Temp)</td>
			<td>2</td>
			<td>0.088</td>
			<td>0.9159</td>
		</tr>
	</tbody>
</table>
Table 2: Analysis of variance(ANOVA). Analysis of variance testing whether changes in life history traits and population growth rate of the resurrected (sub)populations exposed to warming are explained by evolutionary adaptation (populations), plasticity (temperature treatment), and their interaction term (evolution of plasticity). Significant p-values (p&#60;0.05) are shown in bold.
Supplementary Video 1: Sampling of sediment cores. The use of a Big Ben corer is shown. Big Ben is a core tube of approximately 1.5 m in length with an internal tube diameter of 14 cm. It consists of a piston on a rope and a corer head, to which rods are attached to drive the tube into the sediment. A core catcher is used to support the core tube that is deployed from a small vessel. The piston is pushed down into the sediment by gravitational pressure. A framework is used to support the core tube during the extrusion process carried out using a modified bottle jack that pushes the piston upwards. Each sediment layer is collected on a flat metal surface and transferred to transparent sampling bags for long term storage [dark and cold (4 &#176;C) conditions]. <a href="http://cloudflare.jove.com/files/ftp_upload/56637/Supplementary_video1_Sampling.mp4">Please click here to download this file.</a>
Supplementary Video 2: Sediment sieving. The equipment required for sieving sediment is a precision scale, white sampling trays and geological sieves. From each sediment layer, at least 5 g are retained for radiometric dating. The remainder of the sediment is used to isolate ephippia. The sediment is sieved through two geological sieves, one with 1 mm and a second with 125 &#181;m mesh size, piled on top of each other. Medium is poured on the 1 mm mesh sieve to separate clay, large invertebrates, and particulate matter. Medium poured on the second sieve with 125 &#181;m mesh separates D. magna ephippia and small particulate matter. Aliquots of sediment are then transferred to a white sampling tray. D. magna ephippia are spotted by eye in the white background tray. Ephippia from each layer are collected in separate Petri dishes. <a href="http://cloudflare.jove.com/files/ftp_upload/56637/Supplementary_video2_Sieving_sediment.mp4">Please click here to download this file.</a>
Supplementary Video 3: Decapsulation. Under a stereomicroscope, D. magna ephippia are opened with microdissection forceps by applying pressure on the spine of the chitin case. The inner egg membrane is delicately removed and resting eggs gently transferred with a Pasteur pipette to a Petri dish containing 10 mL of medium. <a href="http://cloudflare.jove.com/files/ftp_upload/56637/Supplementary_video3_Decapsulation.mp4">Please click here to download this file.</a>
Supplementary Video 4: Hatching. After exposure to a long photoperiod and 20 &#176;C, embryo development resumes between 48 h and few weeks. When development is complete, the embryos break free from the egg shell and freely swim in the medium. <a href="http://cloudflare.jove.com/files/ftp_upload/56637/Supplementary_video4_Hatching.mp4">Please click here to download this file.</a>

Watch this Scientific Journal Video about Resurrection of Dormant Daphnia magna: Protocol and Applications at JoVE.com

Resurrection of Dormant Daphnia magna: Protocol and Applications

Long-term studies are essential to understanding the process of evolution and the mechanisms of adaptation. Generally, these studies require commitments beyond the life-time of researchers. Here, a powerful method is described that dramatically advances state-of-the-art data collection to generate longitudinal data in natural systems.

Resurrection of Dormant Daphnia magna: Protocol and Applications

Long-term studies enable the identification of eco-evolutionary processes that occur over extended time periods. In addition, they provide key empirical ...

resurrection-of-dormant-daphnia-magna-protocol-and-applications

Universidad de Cartagena UDEC

Research

JoVE Journal

Environment

18.2K Views.  University of Birmingham. Long-term studies are essential to understanding the process of evolution and the mechanisms of adaptation. Generally, these studies require commitments beyond the life-time of researchers. Here, a powerful method is described that dramatically advances state-of-the-art data collection to generate longitudinal data in natural systems.

Video: Resurrection of Dormant Daphnia magna: Protocol and Applications

A Simple Approach to Manipulate Dissolved Oxygen for Animal Behavior Observations

The ability to manipulate dissolved oxygen (DO) in a laboratory setting has significant application to investigate a number of ecological and organismal behavior questions. The protocol described here provides a simple, reproducible, and controlled method to manipulate DO to study behavioral response in aquatic organisms resulting from hypoxic and anoxic conditions. While performing degasification of water with nitrogen is commonly used in laboratory settings, no explicit method for ecological (aquatic) application exists in the literature, and this protocol is the first to describe a protocol to degasify water to observe organismal response. This technique and protocol were developed for direct application for aquatic macroinvertebrates; however, small fish, amphibians, and other aquatic vertebrates could be easily substituted. It allows for easy manipulation of DO levels ranging from 2 mg/L to 11 mg/L with stability for up to a 5 min animal-observation period. Beyond a 5 min observation period water temperatures began to rise, and at 10 min DO levels became too unstable to maintain. The protocol is scalable to the study organism, reproducible, and reliable, allowing for rapid implementation into introductory teaching labs and high-level research applications. The expected results of this technique should relate dissolved oxygen changes to behavioral responses of organisms.

This article describes a simple and reproducible protocol to manipulate dissolved oxygen conditions in a laboratory setting for animal behavior studies. This protocol may be used in both teaching and research laboratory settings to evaluate organismal response of macroinvertebrates, fishes, or amphibians to changes in dissolved oxygen concentration.

The ability to manipulate dissolved oxygen (DO) in a laboratory setting has significant application to investigate a number of ecological and organismal ...

The Barnacle Balanus improvisus as a Marine Model - Culturing and Gene Expression

Barnacles are marine crustaceans with a sessile adult and free-swimming, planktonic larvae. The barnacle Balanus (Amphibalanus) improvisus is particularly relevant as a model for the studies of osmoregulatory mechanisms because of its extreme tolerance to low salinity. It is also widely used as a model of settling biology, in particular in relation to antifouling research. However, natural seasonal spawning yields an unpredictable supply of cyprid larvae for studies. A protocol for the all-year-round culturing of B. improvisus has been developed and a detailed description of all steps in the production line is outlined (i.e., the establishment of adult cultures on panels, the collection and rearing of barnacle larvae, and the administration of feed for adults and larvae). The description also provides guidance on troubleshooting and discusses critical parameters (e.g., the removal of contamination, the production of high-quality feed, the manpower needed, and the importance of high-quality seawater). Each batch from the culturing system maximally yields roughly 12,000 nauplii and can deliver four batches in a week, so up to almost 50,000 larvae per week can be produced. The method used to culture B. improvisus is, probably, to a large extent also applicable to other marine invertebrates with free-swimminglarvae. Protocols are presented for the dissection of various tissues from adults as well as the production of high-quality RNA for studies on gene expression. It is also described how cultured adults and reared cyprids can be utilized in a wide array of experimental designs for examining gene expression in relation to external factors. The use of cultured barnacles in gene expression is illustrated with studies of possible osmoregulatory roles of Na+/K+ ATPase and aquaporins.

The Barnacle Balanus improvisus as a Marine Model - Culturing and Gene Expression

The barnacle Balanus (Amphibalanus) improvisus is a model for studying osmoregulation and antifouling. However, natural seasonal spawning yields an unpredictable supply of cyprid larvae. Here, a protocol for the all-year-round culturing of B. improvisus is described, including the production of larvae. The use of cultured barnacles in gene expression studies is illustrated.

Barnacles are marine crustaceans with a sessile adult and free-swimming, planktonic larvae. The barnacle Balanus (Amphibalanus) improvisus is particularly ...

Basic Methods for the Study of Reproductive Ecology of Fish in Aquaria

Captive-rearing observations are valuable for revealing aspects of fish behavior and ecology when continuous field investigations are impossible. Here, a series of basic techniques are described to enable observations of the reproductive behavior of a wild-caught gobiid fish, as a model, kept in an aquarium. The method focuses on three steps: collection, transport, and observations of reproductive ecology of a substrate spawner. Essential aspects of live fish collection and transport are (1) preventing injury to the fish, and (2) careful acclimation to the aquarium. Preventing harm through injuries such as scratches or a sudden change of water pressure is imperative when collecting live fish, as any physical damage is likely to negatively affect the survival and later behavior of the fish. Careful acclimation to aquaria decreases the incidence death and mitigates the shock of transport. Observations during captive rearing include (1) the identification of individual fish and (2) monitoring spawned eggs without negative effects to the fish or eggs, thereby enabling detailed investigation of the study species' reproductive ecology. The subcutaneous injection of a visible implant elastomer (VIE) tag is a precise method for the subsequent identification of individual fish, and it can be used with a wide size range of fish, with minimal influence on their survival and behavior. If the study species is a substrate spawner that deposits adhesive eggs, an artificial nest site constructed from polyvinyl chloride (PVC) pipe with the addition of a removable waterproof sheet will facilitate counting and monitoring the eggs, lessening the investigator's influence on the nest-holding and egg-guarding behavior of the fish. Although this basic method entails techniques that are seldom mentioned in detail in research articles, they are fundamental for undertaking experiments that require the captive rearing of a wild fish.

A series of basic methods to enable the study of the reproductive ecology of fish kept in aquaria are described. These are useful protocols for collecting fish using SCUBA, transporting live fish, and observing the reproductive behavior of wild-caught fish kept in aquaria.

Captive-rearing observations are valuable for revealing aspects of fish behavior and ecology when continuous field investigations are impossible. Here, a ...

Behavior

Cryopreservation of PGCs for Long-Term Preservation of Drosophila Strains

Primordial Germ Cell Cryopreservation and Revival of Drosophila Strains

Drosophila strains must be maintained by the frequent transfer of adult flies to new vials. This carries a danger of mutational deterioration and phenotypic changes. Development of an alternative method for long-term preservation without such changes is therefore imperative. Despite previous successful attempts, cryopreservation of Drosophila embryos is still not of practical use because of low reproducibility. Here, we describe a protocol for primordial germ cell (PGC) cryopreservation and strain revival via transplantation of cryopreserved PGCs into agametic Drosophila melanogaster (D. melanogaster) host embryos. PGCs are highly permeable to cryoprotective agents (CPAs), and developmental and morphological variation among strains is less problematic than in embryo cryopreservation. In this method, PGCs are collected from approximately 30 donor embryos, loaded into a needle after CPA treatment, and then cryopreserved in liquid nitrogen. To produce donor-derived gametes, the cryopreserved PGCs in a needle are thawed and then deposited into approximately 15&#160;agametic host embryos. A frequency of at least 15% fertile flies was achieved with this protocol, and the number of progeny per fertile couple was always more than enough to revive the original strain (the average progeny number being 77.2 &#177; 7.1), indicating the ability of cryopreserved PGCs to become germline stem cells. The average number of fertile flies per needle was 1.1 &#177; 0.2, and 9 out of 26 needles produced two or more fertile progeny. It was found that 11 needles are enough to produce 6 or more progeny, in which at least one female and one male are likely included. The agametic host makes it possible to revive the strain quickly by simply crossing newly emerged female and male flies. In addition, PGCs have the potential to be used in genetic engineering applications, such as genome editing.

Author Spotlight: PGC Cryopreservation &#8211; An Alternative Approach Towards Long-Term Preservation of Drosophila Strains 

A long-term preservation method for Drosophila strains as an alternative to the frequent transfer of adult flies to fresh food vials is highly desirable. This protocol describes the cryopreservation of Drosophila primordial germ cells and strain revival via their transplantation to agametic host embryos.

Drosophila strains must be maintained by the frequent transfer of adult flies to new vials. This carries a danger of mutational deterioration and ...

Genetics

In vivo Imaging of Intact Drosophila Larvae at Sub-cellular Resolution

Recent improvements in optical imaging, genetically encoded fluorophores and genetic tools allowing efficient establishment of desired transgenic animal lines have enabled biological processes to be studied in the context of a living, and in some instances even behaving, organism. In this protocol we will describe how to anesthetize intact Drosophila larvae, using the volatile anesthetic desflurane, to follow the development and plasticity of synaptic populations at sub-cellular resolution1-3. While other useful methods to anesthetize Drosophila melanogaster larvae have been previously described4,5,6,7,8, the protocol presented herein demonstrates significant improvements due to the following combined key features: (1) A very high degree of anesthetization; even the heart beat is arrested allowing for lateral resolution of up to 150 nm1, (2) a high survival rate of &gt; 90% per anesthetization cycle, permitting the recording of more than five time-points over a period of hours to days2 and (3) a high sensitivity enabling us in 2 instances to study the dynamics of proteins expressed at physiological levels. In detail, we were able to visualize the postsynaptic glutamate receptor subunit GluR-IIA expressed via the endogenous promoter1 in stable transgenic lines and the exon trap line FasII-GFP1. (4) In contrast to other methods4,7 the larvae can be imaged not only alive, but also intact (i.e. non-dissected) allowing observation to occur over a number of days1. The accompanying video details the function of individual parts of the in vivo imaging chamber2,3, the correct mounting of the larvae, the anesthetization procedure, how to re-identify specific positions within a larva and the safe removal of the larvae from the imaging chamber.

In vivo Imaging of Intact Drosophila Larvae at Sub-cellular Resolution

This protocol describes a reliable method for anesthetization and imaging of intact Drosophila melanogaster larvae. We have utilized the volatile anesthetic desflurane to allow for repetitive imaging at sub-cellular resolution and re-identification of structures for up to a few days1.

Recent improvements in optical imaging, genetically encoded fluorophores and genetic tools allowing efficient establishment of desired transgenic animal ...

Biology

Pharmacological and Functional Genetic Assays to Manipulate Regeneration of the Planarian Dugesia japonica

Free-living planarian flatworms have a long history of experimental usage owing to their remarkable regenerative abilities1. Small fragments excised from these animals reform the original body plan following regeneration of missing body structures. For example if a 'trunk' fragment is cut from an intact worm, a new 'head' will regenerate anteriorly and a 'tail' will regenerate posteriorly restoring the original 'head-to-tail' polarity of body structures prior to amputation (Figure 1A).
Regeneration is driven by planarian stem cells, known as 'neoblasts' which differentiate into ~30 different cell types during normal body homeostasis and enforced tissue regeneration. This regenerative process is robust and easy to demonstrate. Owing to the dedication of several pioneering labs, many tools and functional genetic methods have now been optimized for this model system. Consequently, considerable recent progress has been made in understanding and manipulating the molecular events underpinning planarian developmental plasticity2-9.
The planarian model system will be of interest to a broad range of scientists. For neuroscientists, the model affords the opportunity to study the regeneration of an entire nervous system, rather than simply the regrowth/repair of single nerve cell process that typically are the focus of study in many established models. Planarians express a plethora of neurotransmitters10, represent an important system for studying evolution of the central nervous system11, 12 and have behavioral screening potential13, 14. 
Regenerative outcomes are amenable to manipulation by pharmacological and genetic apparoaches. For example, drugs can be screened for effects on regeneration simply by placing body fragments in drug-containing solutions at different time points after amputation. The role of individual genes can be studied using knockdown methods (in vivo RNAi), which can be achieved either through cycles of microinjection or by feeding bacterially-expressed dsRNA constructs8, 9, 15. Both approaches can produce visually striking phenotypes at high penetrance- for example, regeneration of bipolar animals16-21. To facilitate adoption of this model and implementation of such methods, we showcase in this video article protocols for pharmacological and genetic assays (in vivo RNAi by feeding) using the planarian Dugesia japonica.

Pharmacological and Functional Genetic Assays to Manipulate Regeneration of the Planarian Dugesia japonica

An attractive model for studying stem cell differentiation within a live animal is the planarian flatworm. Regeneration is studied by simple amputation experiments that are easily performed in a basic laboratory and are amenable to pharmacological and genetic (in vivo RNAi) manipulation as detailed by protocols in this article.

Free-living planarian flatworms have a long history of experimental usage owing to their remarkable regenerative abilities1. Small fragments excised from ...

A Method of Permeabilization of Drosophila Embryos for Assays of Small Molecule Activity

The Drosophila embryo has long been a powerful laboratory model for elucidating molecular and genetic mechanisms that control development. The ease of genetic manipulations with this model has supplanted pharmacological approaches that are commonplace in other animal models and cell-based assays. Here we describe recent advances in a protocol that enables application of small molecules to the developing fruit fly embryo. The method details steps to overcome the impermeability of the eggshell while maintaining embryo viability. Eggshell permeabilization across a broad range of developmental stages is achieved by application of a previously described d-limonene embryo permeabilization solvent (EPS1) and by aging embryos at reduced temperature (18 &#176;C) prior to treatments. In addition, use of a far-red dye (CY5) as a permeabilization indicator is described, which is compatible with downstream applications involving standard red and green fluorescent dyes in live and fixed preparations. This protocol is applicable to studies using bioactive compounds to probe developmental mechanisms as well as for studies aimed at evaluating teratogenic or pharmacologic activity of uncharacterized small molecules.

A Method of Permeabilization of Drosophila Embryos for Assays of Small Molecule Activity

Introduction of small molecules to the developing Drosophila embryo offers great potential for characterizing biological activity of novel compounds, drugs, and toxins as well as for probing fundamental developmental pathways. Methods described herein outline steps that overcome natural barriers to this approach, expanding the utility of the Drosophila embryo model.

The Drosophila embryo has long been a powerful laboratory model for elucidating molecular and genetic mechanisms that control development. The ease of ...

Neural Stem Cell Reactivation in Cultured Drosophila Brain Explants

Neural stem cells (NSCs) have the ability to proliferate, differentiate, undergo apoptosis, and even enter and exit quiescence. Many of these processes are controlled by the complex interplay between NSC intrinsic genetic programs with NSC extrinsic factors, local and systemic. In the genetic model organism, Drosophila melanogaster, NSCs, known as neuroblasts (NBs), switch from quiescence to proliferation during the embryonic to larval transition. During this time, larvae emerge from their eggshells and begin crawling, seeking out dietary nutrients. In response to animal feeding, the fat body, an endocrine organ with lipid storage capacity, produces a signal, which is released systemically into the circulating hemolymph. In response to the fat body-derived signal (FBDS), Drosophila insulin-like peptides (Dilps) are produced and released from brain neurosecretory neurons and glia, leading to downstream activation of PI3-kinase growth signaling in NBs and their glial and tracheal niche. Although this is the current model for how NBs switch from quiescence to proliferation, the nature of the FBDS extrinsic cue remains elusive. To better understand how NB extrinsic systemic cues regulate exit from quiescence, a method was developed to culture early larval brains in vitro before animal feeding. With this method, exogenous factors can be supplied to the culture media and NB exit from quiescence assayed. We found that exogenous insulin is sufficient to reactivate NBs from quiescence in whole-brain explants. Because this method is well-suited for large-scale screens, we aim to identify additional extrinsic cues that regulate NB quiescence versus proliferation decisions. Because the genes and pathways that regulate NSC proliferation decisions are evolutionarily conserved, results from this assay could provide insight into improving regenerative therapies in the clinic.

Neural Stem Cell Reactivation in Cultured Drosophila Brain Explants

A method to reactivate quiescent neural stem cells in cultured Drosophila brain explants has been established. Using this method, the role of systemic signals can be uncoupled from tissue-intrinsic signals in the regulation of neural stem cell quiescence, entry and exit.

Neural stem cells (NSCs) have the ability to proliferate, differentiate, undergo apoptosis, and even enter and exit quiescence. Many of these processes ...

Neuroscience

Reactivation of Demembranated Cell Models in Chlamydomonas reinhardtii

Since the historical experiment on the contraction of glycerinated muscle by adding ATP, which Szent-Gy&#246;rgyi demonstrated in the mid-20th century, in vitro reactivation of demembranated cells has been a traditional and potent way to examine cell motility. The fundamental advantage of this experimental method is that the composition of the reactivation solution may be easily changed. For example, a high-Ca2+ concentration environment that occurs only temporarily due to membrane excitation in vivo can be replicated in the lab. Eukaryotic cilia (a.k.a. flagella) are elaborate motility machinery whose regulatory mechanisms are still to be clarified. The unicellular green alga Chlamydomonas reinhardtii is an excellent model organism in the research field of cilia. The reactivation experiments using demembranated cell models of C. reinhardtii and their derivatives, such as demembranated axonemes of isolated cilia, have significantly contributed to understanding the molecular mechanisms of ciliary motility. Those experiments clarified that ATP energizes ciliary motility and that various cellular signals, including Ca2+, cAMP, and reactive oxygen species, modulate ciliary movements. The precise method for demembranation of C. reinhardtii cells and reactivation of the cell models is described here.

Reactivation of Demembranated Cell Models in Chlamydomonas reinhardtii

The in vitro reactivation of motile cells is a crucial experiment in understanding the mechanisms of cell motility. The protocol describes reactivating the demembranated cell models of Chlamydomonas reinhardtii, a model organism to study cilia/flagella.

Since the historical experiment on the contraction of glycerinated muscle by adding ATP, which Szent-Gy&#246;rgyi demonstrated in the mid-20th century, in ...

Investigating Collective Behaviors in Nematodes Using Dog Food Agar Medium

The Large-Scale Cultivation of Nematodes to Study Their Collective Behaviors

Animals exhibit dynamic collective behaviors, as observed in flocks of birds, schools of fish, and crowds of humans. The collective behaviors of animals have been investigated in the fields of both biology and physics. In the laboratory, researchers have used various model animals such as the fruit fly and zebrafish for approximately a century, but it has remained a major challenge to study large-scale complex collective behavior orchestrated by these genetically tractable model animals. This paper presents a protocol to create an experimental system of collective behaviors in Caenorhabditis elegans. The propagated worms climb on the lid of the Petri plate and show collective swarming behavior. The system also controls worm interactions and behaviors by changing the humidity and light stimulation. This system allows us to examine the mechanisms underlying collective behaviors by changing environmental conditions and examining the effects of individual-level locomotion on collective behaviors using mutants. Thus, the system is useful for future research in both physics and biology.

Author Spotlight: Collective Behavioral Analysis of the Nematode, Caenorhabditis elegans

Here, a system is reported for studying the collective behaviors of nematodes by culturing them in bulk using dog food agar medium. This system allows researchers to propagate large numbers of dauer worms and can be applied to Caenorhabditis elegans and other related species.

Animals exhibit dynamic collective behaviors, as observed in flocks of birds, schools of fish, and crowds of humans. The collective behaviors of animals ...

Resurrection of Dormant Daphnia magna: Protocol and Applications

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A Method of Permeabilization of Drosophila Embryos for Assays of Small Molecule Activity

Neural Stem Cell Reactivation in Cultured Drosophila Brain Explants

Reactivation of Demembranated Cell Models in Chlamydomonas reinhardtii

The Large-Scale Cultivation of Nematodes to Study Their Collective Behaviors