A typical day spent in our lab’s aquarium room will find me soaked, top-to-bottom, in seawater. The other members of my lab seem not to have this issue, but I’ve always seemed to get “in” to my science, literally, when it comes to these types of chores. The trouble with marine models is that they require salt water, and lots of it. This requires that I balance half of my small frame over the edge of a 44 gallon drum so that I may reach to stir in any last bits of un-dissolved salt. I pipette a small drop of the freshly made salt water onto the spectrometer, closing one eye as I bring the device to the other, pointing it up to the light in order to check that the salinity is within 28 – 32 PPT. This always makes me feel like a pirate. Normally, the day-to-day care of the stock tanks is hurried time away from research, but on the days of failed experiments, these mundane tasks—making seawater, cleaning tanks, fixing pumps, and the like—can be a satisfying reprieve. Until, that is, you get a giant mouthful of dirty seawater while attempting to start the flow of an aquarium siphon to vacuum the bottom of the tanks.

When I’m not maintaining the tanks in the pleasantly warm and humid aquarium room, I can be found in the back corner of the lab, carefully manipulating delicate embryos under the microscope with homemade tools; at my bench, changing wash after wash of an immunofluorescent antibody stain; or in the tiny windowless room filled predominantly by the very expensive confocal microscope, watching the attached computer screen hopefully as bright layers of color appear, digital slice by digital slice. The images that form are the visualized results of weeks of work. Until you actually see your results, anything is possible, claimed one past mentor; I’m not sure how much I buy into this “Schrödinger’s results” mindset, but I’m sometimes afraid to look just the same.

A new model

Par … hi … allie? The name of this up-and-coming model organism doesn’t quite roll off the tongue, and I have yet to meet someone who can pronounce Parhyale for the first time without stumbling. Parhyale hawaiensis, perhaps better known as “beach hoppers” or “sand fleas”, are amphipod crustaceans that like to hang out on decaying plant matter in intertidal, equatorial waters worldwide, or, in our lab, on carrots and air bubblers.

If you haven’t had the chance to meet (or mispronounce) this up-and-coming model organism, don’t worry—you will, because these guys are making quite the splash in the field of developmental biology. When I first joined Parhyale pioneer Nipam Patel’s lab as an incoming graduate student at UC Berkeley, I knew nothing of them. I came from a vertebrate-centric background, and had little experience with arthropods (or anything not warm and fuzzy, for that matter). So green was I to the frequently cited arthropod orders, that when I first heard the term “amphipod” I thought I was mishearing the term “arthropod”. Parhyale were my first introduction to this new and wonderful world of invertebrates, and I would soon learn just how important their appendages are for answering questions about morphological diversity.  Indeed, the phylum Arthropoda means “jointed foot” and amphipod refers to the two different orientations of the thoracic appendages in this order.

Using Parhyale, our lab studies developmental mechanisms that pattern the body plan and the evolutionary changes that have led to the diversity of arthropod appendages. With their amiability to laboratory manipulation, total cleavage forming stereotyped lineages early in development, and boundless supply of embryos, Parhyale make an excellent model for developmental studies.

Tupperwares of love

The first thing you may notice as you enter our lab is the hum of the air bubblers from the Parhyale tanks—clusters of various sized Tupperware that populate each bench. The tanks soon fall into the background drone of freezers and other electronic equipment. The tanks vary in cleanliness (this may or may not be correlated to the availability of undergrads), though the Parhyale don’t seem to mind. They are detritivores, and live off of decaying organic matter in the ocean. In the lab, we feed them carrots. Yes, carrots. They cling to them in droves; presumably eating whatever it is that grows on them.

Indeed, that’s what Parhyale were chosen for—their hardiness. When Bill Browne (then a graduate student in the Patel lab) went to Chicago’s Shedd Aquarium in search of new crustaceans for the lab, he didn’t go to the display tanks. Rather, he surveyed the filtration system and discovered an ideal lab animal—one that survives on garbage without the need for constant care. This hardiness can be attributed to the constant, naturally occurring temperature and salinity fluctuations of the shallow water and intertidal habitats they have evolved to flourish.

But Parhyale offers more than just hardiness and ease of bench top maintenance. They offer fecundity; it’s part of their charm. “Aw, look at all of the mommies swimming around paired with their babies!” exclaimed one colleague when I first showed her my research subjects. No, I had to explain, when a boy Parhyale loves a girl Parhyale, the larger male shows his affections by dragging her around with him in pre-mating “amplexus” until she molts and he fertilizes the eggs and releases her. Thus begins a new cycle of embryo production. Development labs are necessarily centered to the reproductive agenda of their animals, but there is no finickiness—or shortages of gravid females—in our Tupperwares of love. Females can produce embryos every two weeks after (quickly) reaching sexual maturity, and, as tropical species tend to do, they produce broods year round. To learn more about these guys, check out The crustacean Parhyale hawaiensis: A New Model for Arthropod Development.

Parhyale Amplexus. Drawing by Jessica Poon.

A day in the life of a Parhyale wrangler

On any given day, my tanks contain a good mix of males, females, mate pairs, and gravid females. But I am after something specific—one- and two-cell embryos. To increase my chances of catching this four to nine hour window of a 10-day embryogenesis, I go for the pairs. I cut the end off of a 3 mL transfer pipette so that it’s wide enough (barely) to suck up a mating pair, and, as I eye my quarry among the carrots, pH-buffering gravel, and non-paired individuals, let the rodeo begin. I’m good at this now; people new to the lab are impressed with the quickness at which I can pluck my chosen paired Parhyale out of the water, despite their fast reactions and expert swimming. These pairs are placed in a separate tank and regularly observed for detachment. Newly single males are removed, and the freshly gravid females are detained for embryo collection; adding a few microliters of clove oil to their water will knock them out.

Parhyale, like all amphipods, carry their embryos in a brood pouch on their ventral side. Delicately clasping the sleeping body with forceps, I use the smooth end of a glass pipette pulled and rounded over a flame to gently scrape along her ventral pouch; the embryos pop up and drift away. Done correctly, embryo collection is fairly non-invasive for the female. The clove oil anesthetic soon wears off and the females can be returned to their tanks to resume their breeding cycle.

The one and two-cell embryos are prime for microinjection. siRNA against my gene of choice is loaded into a microinjection needle pulled from a small capillary tube. The loud “WHAP” made by the force of the needle puller separating as a pulled capillary tube breaks in two still makes me jump. My dedicated undergraduate, Jennifer Wang, does the lion’s share of the siRNA injections that will knockdown the targeted mRNA.  If we want to knock down expression throughout the entire animal, we inject the one-cell embryo, or both cells at the two-cell stage.  For some experiments, we can also achieve lineage specific knockdown by injecting one or more cells at the eight-cell stage.

Injection of a single blastomere of an eight cell embryo.

Fate map of the eight cell stage of Parhyale development.

Once injected, this is when we have to really care for them. Parhyale embryos will develop just fine in a tissue culture dish, but the water must be changed several times a week to avoid fungal contamination and death. It’s easy to keep track of embryonic stages in Parhyale thanks to the detailed Parhyale staging system of Browne, et al. There are 30 stages divided by time point and visual characteristics, many of which have lab nicknames names such as “frosted-soccer ball” stage or simply “leggy”, which is the stage I’m after.

Now comes the fun part, the part where I sit for hours dissecting out embryo after embryo from their extra-embryonic membranes. Learning to dissect out an embryo—intact—requires countless hours of frustration at the beginning, but it’s rewarding, and, eventually (dare I say)—fun, once you get the hang of it. To accomplish these tiny dissections, we thread small pieces of tungsten wire into insulin needles, and sharpen these dissection needles by dipping their ends into a beaker of sodium hydroxide with a current running through it. I use these tools to secure a rolling embryo, poking a hole through both membranes to allow yolk to flow out, and the fixation solution (4% paraformaldehyde in sea water) to flow in. Teasing the embryo, the outer membrane seems to pop right off. Removing the inner membrane takes much more careful manipulation, and much more patience.

Once I have a tube full of fixed embryos with their membranes removed, I use immunofluorescent antibody stains to visualize gene expression patterns and structures within the embryos. Easy as making cookies—except that my antibody stains usually turn out more successfully than my half-hearted attempts at baking (turns out that antibody incubation times are much more flexible than bake times). The tricky part is not accidently sucking up a small (sometimes floating) embryo while performing wash after wash in PBS with Triton. The last step is clearing with glycerol; the embryos are now ready to be mounted. I mount one Parhyale embryo per slide, carefully splaying out each limb and using the twisted edge of a Kimwipe to suck up any excess glycerol. I hold my breath as I place the cover slip over a particularly nice mount—it’s always the perfect ones that seem to get messed up during this step. If I am lucky, I can snag an opening on the confocal; we share the microscope with the entire floor, and scheduling can be tight. I’m in luck, there’s an evening slot. With no one behind me, I’m able to confocal late into the night. As the confocal scans, the gene expression patterns are slowly revealed, ventral to dorsal, in a satisfying mix of my chosen colors: aqua, neon green, Christmas red, and magenta bordering on pink.

Immunofluorescent stains of Parhyale embryos.
Left: Blue = DAPI, Magenta = anti-HRP (nervous system).  Center: Red = Engrailed, Green = Ultrabithorax.  Right: Images combined

Right now I am focused on the function of homeotic genes in crustaceans, and how evolutionary changes in these genes have contributed to the diversification of arthropod appendages.  I use siRNAs to knock down individual Hox genes, and then use various markers to examine changes in the ectoderm, nervous system, and musculature to see if all parts of the segment are transformed to understand how the fate of these different structures is coordinated during development.  As I make my way through my 3^rd year as a graduate student, I now consider myself one of the few, the proud, the Parhyale researchers. My friends (the non-sciency ones) consider me unemployed.

This post is part of a series on a day in the life of developmental biology labs working on different model organisms. You can read the introduction to the series here and read other posts in this series here.

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We realise that a lot of the people who register for an account with the Node are not experienced bloggers. It can be hard to write your first blog post: What should I write about? What is the best style? How long should a post be?

To help you write your first post (or make it easier to write more), we created a list of writing tips for Node bloggers. Have a read through and let us know what you think!

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Image: Wellcome Library, London

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Those lucky scientists, who study organisms which allow live imaging experiments to be effectively performed, do not always appreciate what a luxury it is to watch the tissue of interest develop in time. Scientists who work on less accessible models that take long time to develop to term, look with envy on the beautiful movies of other model systems that show the movement and interaction of the cells, as well as the actions and reactions in the vicinity of a region of interest. Although he developing wing disc in the drosophila larvae serves as a long standing model to study the coordination between tissue growth and patterning, it is not an easy structure to watch as it develops, since it its increase of roughly thousand fold in cell number occurs in a timespan of five days. Larval movement and muscle contractions associated with feeding and other processes  make it impossible to image the entire process in vivo and in vitro culture of isolated imaginal discs during the  entire growth has not been possible thus far.

In contrast to the limitations with regard to live imaging, the wing disc does serve as an excellent genetic model system. Moreover, since the wing is dispensable for the viability of the fly, a large number of elegant genetic screens allowed to put forward a fine blueprint of its development after decades of research from hundreds of scientists. The larval part of the blueprint that we have gained on wing growth is based on dissecting multiple wing discs of different developmental stages, comparing them to each other and merging the information coming from different discs from the same time point. Using this approach, the information that we can acquire from an individual disc is very limited. This is a limitation that is not restricted to the study of the growth of the wing imaginal disc; other developing organs that take long time to develop suffer from this caveat. Our aim in developing Raeppli was to maximise the information we can extract from each sample (Kanca et al. 2014). We envisioned that marking the lineage of each cells of a young wing disc with a fluorescent protein would allow us to distinguish the contribution of each of those lineages to the adult structure.

We got inspired by the Brainbow and related techniques that are most often used to track the projections of neurones in the developing nervous system (Livet et al. 2007, Hampel et al. 2011, Hadjieconomou et al. 2011) . These techniques use mutually exclusive recombination events to select one of three fluorescent proteins, in individual cells, thereby also marking permanently their progeny. Use of multiple such constructs generates an RGB like colour coding of different cells. We started from this basic idea but diverged from it along the implementation of the idea.
 
Cre recombinase, which is used for mutually exclusive recombination in Brainbow, is quite toxic in Drosophila, and rarely used in fly research.  The Flippase recombinase is somewhat inefficient and causes recombination in a fraction of cells only. We thus opted to use the integrase system, since recombination is both irreversible and efficient. The first versions of constructs we cloned coded for six fluorescent proteins in order to maximise the colour choice per construct. Although these constructs in principle worked to mark cells with different colour combinations, the 6 fluorescent proteins could not be spectrally separated efficiently since there was substantial bleed-through between different colour channels. Although this generates different shades of colours, when two copies of the construct are used the interpretation of the data gets complicated. Thus, for a second version of constructs,we decided to use 4 fluorescent proteins that can be spectrally separated from each other in sequential scans. The fluorescent proteins we chose were selected to be bright and to be usable 1) in combination with GFP, which is applied as a widespread marker, and 2) in combination with far red antibodies. This further increases the amount of information that can be acquired from the sample in addition to the lineage information. We generated two different versions of Raeppli, one where all the fluorescent proteins are directed to membrane by Ras Farnasylation sequence (CAAX), thus marking cell membranes, second where all the fluorescent proteins are directed to nucleus by using Nuclear Localisation Signal (NLS).  By using two copies of the construct, we could mark more than 90% of the cells in a wing disc with diverse colour combinations. Moreover, since the fluorescent proteins were bright enough to be visualised through the larval cuticle, we were capable to image the growth of a single wing disc over the three larval periods by acquiring snapshots of the disc from an anaesthetised larva. Multiple individually marked clones served as reference points to compare the different time points. Moreover, we opted to increase the flexibility of usage of Raeppli by including a cassette that, when unrecombined, can respond to either Gal4 or LexA, and when recombined by Cre, can respond to either one or the other binary system. Thus, for routine uses one can express the proteins with any driver that one wants, but for more specific uses, such as over-expression studies using an additional transgene, in combination to lineage analysis, one can use one binary system for detection of Raeppli and the other for manipulating the cells.

Although the analyses shown in the paper are confined the study of wild type tissue behaviour, we are planning to explore  the flexibility and efficacy of Raeppli and analyse the whole tissue response induced by perturbations in parts of the tissue. Due to the flexibility of Raeppli, we believe that its use will help to address numerous interesting scientific questions, both at the cellular as well as at the tissue level. Finally, together with recent advances in cell labeling tools, Raeppli will help to shed new light on organogenesis (Boulina et al. 2013, Wortley et al. 2013, Kanca et al. 2014) .

References:

1.    Boulina, M., Samarajeewa, H., Baker, J. D., Kim, M. D. & Chiba, A. Live imaging of multicolor-labeled cells in Drosophila. Development 140, 1605–1613 (2013).

2.    Hadjieconomou, D. et al. Flybow: genetic multicolor cell labeling for neural circuit analysis in Drosophila melanogaster. Nat. Methods 8, 260–266 (2011).

3.    Hampel, S. et al. Drosophila Brainbow: a recombinase-based fluorescence labeling technique to subdivide neural expression patterns. Nat. Methods 8, 253–259 (2011).

4.    Kanca, O., Caussinus, E., Denes, A. S., Percival-Smith, A. & Affolter, M. Raeppli: a whole-tissue labeling tool for live imaging of Drosophila development. Development 141, 472–480 (2014).

5.    Livet, J. et al. Transgenic strategies for combinatorial expression of fluorescent proteins in the nervous system. Nature 450, 56–62 (2007).

6.    Worley, M. I., Setiawan, L. & Hariharan, I. K. TIE-DYE: a combinatorial marking system to visualize and genetically manipulate clones during development in Drosophila melanogaster. Development 140, 3275–3284 (2013).

(9 votes)

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Here are the highlights from the current issue of Development:

Hoxb1b gets the neural tube into shape

Hox genes are classically known for their roles in patterning the anterior-posterior axis of animals. Now, on p. 639, Mihaela Žigman, Cecilia Moens and colleagues uncover a new role for Hoxb1b in regulating cell shape, oriented divisions and microtubule dynamics in the developing zebrafish neural tube. The researchers first identify a zebrafish mutant that carries a point mutation in hoxb1b, a gene that shares ancestral functions with mammalian Hoxa1. These mutants, they report, exhibit classical homeotic transformations associated with Hoxa1 mutations in mice. Unexpectedly, however, these mutants also show defective neuroepithelial morphogenesis within the developing hindbrain neural tube. The researchers further show that the hoxb1b mutation does not affect apico-basal or adherens junction-based polarisation, nor the proliferation or differentiation rate of neural progenitors. Instead, Hoxb1b regulates mitotic spindle orientation and the shape of progenitor cells. This function is linked to a cell-non-autonomous role for Hoxb1b in regulating microtubule dynamics. The authors thus propose that, by regulating microtubule dynamics and cell shape, Hox genes can influence global tissue morphogenetic events.

A new model for bivalency

Histone H3 lysine 4 trimethylation (H3K4me3) is a universal epigenetic mark that is deposited by histone methyltransferases. This mark can be found in the context of bivalent promoters, which harbour both repressive H3K4me3 and active H3K27me3 marks and hence are thought to be poised for lineage-specific activation or repression. Here, Francis Stewart, Henk Stunnenberg and co-workers challenge this model of poising (p. 526). They first show that the H3K4 methyltransferase Mll2 is responsible for H3K4me3 on bivalent promoters in embryonic stem cells (ESCs). Accordingly, the researchers find that Mll2 is bound to bivalent promoters but also to active promoters. By contrast, another H3K4 methyltransferase, Set1C, is bound to active but not bivalent promoters. Importantly, they observe that Mll2-deficent ESCs, which lack H3K4me3 on bivalent promoters, exhibit normal transcription responsiveness, thus arguing against a model of poising. Based on these and other findings, the authors propose that Mll2 acts as a pioneer methyltransferase and that Polygroup group action on bivalent promoters blocks the establishment of active Set1C-bound promoters.

InSpired dendrite architecture

The correct architecture of dendritic trees is essential for the wiring and function of neuronal circuits. A number of cell extrinsic factors are known to regulate dendrite shape and patterning, but here Don van Meyel and co-workers show that the transcription factor Longitudinals Lacking (Lola) regulates expression of the actin nucleation protein Spire (Spir) to sculpt dendrite architecture in Drosophila (p. 650). The researchers show that Lola is expressed in dendritic arborisation (da) neurons of the Drosophila peripheral nervous system. They further demonstrate that Lola controls the number, growth and distribution of dendrite branches in da neurons. Loss of Lola also leads to increased expression of Spir, which in turn causes increased formation of abnormal and inappropriately positioned actin-rich branches. In line with this, the authors report that Spir promotes F-actin nucleation and regulates dendrite positioning. Together, these findings suggest that Lola acts to limit the expression of Spir within da neurons, thus ensuring balanced control of the actin cytoskeleton and regulated dendrite morphogenesis.

miR-335 shapes an endoderm transcription factor gradient

Morphogen and transcription factor gradients are known to pattern tissues during development, but how these gradients are established is unclear. Using mouse embryos, embryonic stem cells (ESCs) and mathematical modelling, Heiko Lickert and colleagues show that the microRNA miR-335 fine-tunes a transcription factor gradient in the endoderm (p. 514). The researchers identify miR-335 as a microRNA that is differentially regulated during mesendoderm differentiation. They further show that miR-335 is expressed and functions transiently in endoderm progenitors and later during mesoderm formation. Importantly, miR-335 targets mRNAs encoding the endoderm-determining transcription factors Foxa2 and Sox17; miR-335 overexpression blocks endoderm differentiation in ESCs and, conversely, inhibition of miR-335 activity leads to Foxa2 and Sox17 accumulation and increased endoderm formation. Finally, mathematical modelling incorporating both microRNA and protein turnover rates predicts that miR-335 can shape a gradient of Foxa2 and Sox17 in the endoderm, and this prediction is confirmed experimentally. Overall, these findings highlight that a microRNA can shape a transcription factor gradient in time and space.

Eyeing up nutrient control of stem and progenitor cells

It is known that nutrient availability affects cell proliferation, but how nutrients affect the proliferation-differentiation programme of cells is unclear. On p. 697, Nicola Love and colleagues address this issue, using the ciliary marginal zone (CMZ) of the Xenopus retina as a model. They find that nutrient deprivation (ND) reduces the proliferation, and hence the number, of committed retinal progenitors in the CMZ. By contrast, retinal stem cells at the CMZ peripheral edge are relatively insensitive to ND. Furthermore, ND prevents cells from acquiring a committed progenitor fate, suggesting the presence of a nutrient-sensitive restriction point in the retinal progenitor proliferation-differentiation programme. Finally, the authors show that this restriction point involves mTOR signalling; blocking mTOR mimics many of the effects of ND, whereas activation of mTOR stimulates differentiation. Together, these findings suggest that an mTOR-dependent restriction point in the proliferation-differentiation programme of retinal progenitors exists to couple nutrient availability to tissue growth and development, thus allowing regrowth in ND tissue when conditions of plenty return.

Jaw-dropping differences in the neural crest

Variation in jaw size has been crucial to the evolution and adaptation of vertebrates. On p. 674, Jennifer Fish, Richard Schneider and colleagues explore the mechanisms by which duck and quail achieve distinct jaw sizes, testing the hypothesis that differences in neural crest (NC) biology contribute to species-specific differences in jaw size. The researchers show that the total sizes of the pre-migratory NC progenitor populations in duck and quail are similar. However, the midbrain region, which generates jaw NC precursors, is wider and shorter in duck owing to an anterior shift in brain regionalisation. Furthermore, they report, more pre-migratory NC precursors are allocated to the midbrain in duck, which gives rise to an increased number of post-migratory NC cells within the duck mandibular arch. Finally, differences in proliferation lead to an increase in the size of the duck mandibular arch relative to that of the quail. Thus, the authors propose, the larger jaw size of duck is the result of at least three distinct developmental events.

PLUS…

How to make spinal motor neurons

All muscle movements, including breathing, walking, and fine motor skills rely on the function of spinal motor neurons. Here, Kevin Eggan and colleagues discuss how the logic of spinal motor neuron development has been applied to generate motor neurons either from pluripotent stem cells by directed differentiation and transcriptional programming, or from somatic cells by direct lineage conversion. See the Primer on p. 491

Lung development: orchestrating the generation and regeneration of a complex organ

The respiratory system, which consists of the lungs, trachea and associated vasculature, is essential for terrestrial life. In recent years, extensive progress has been made in defining the temporal progression of lung development.  This has led to exciting discoveries including the derivation of lung epithelium from stem cells and the discovery of new targets for therapeutics. Michael Herriges and Ed Morrisey highlight review these recent advances in our understanding of lung development and regeneration. See the Review on p. 502

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Here is our monthly round-up of some of the interesting content that we spotted around the internet:

News & Research:

– Nature revealed their list of 10 people that mattered in 2013, while the UK Science Council revealed its list of 100 leading UK practicing scientists.

– This Christmas saw developmental biologist Alison Woollard present the Royal Institution Christmas lectures. You can watch them on the Royal Institution website.

– Nature News & Views published a piece on the importance of senescence in embryonic development.

– And stem cells have been in the limelight, with articles featuring updates on the current controversial situation in Italy and Woo Suk Hwang, 10 years after his stem cell cloning fraud scandal.

Weird & Wonderful:

– An Australian science agency apologised to a little girl for their lack of dragon research.

– The ASCB website suggested a few uses for those old conference posters.

– If you are a scientist and a Monty Python fan you might want to check a new hashtag trending on Twitter- #MontyPythonScience.

Beautiful & Interesting images:

– If you are a PhD student you might find this list of gifs displaying 25 painful problems of graduate students (besides their thesis) highly amusing.

– We spotted several beautiful images: colourful neural stem cell images, a delicate tree that is actually a protist and a dividing cell made of fused glass.

– And we found a great photo of how a scientist protests:

What do we want? pic.twitter.com/mwyO4fyoVU

— Dr Brooke Magnanti (@bmagnanti) January 7, 2014

Videos worth watching:

– The Naked Scientists dedicated one of their recent podcasts to developmental biology.

– The Royal Institution listed their favourite science movies of 2013, which includes this beautiful video on the life cycle of sea urchins by Parafilms:

Keep up with this and other content, including all Node posts and deadlines of coming meetings, by following the Node on Twitter.

Alison’s image by Paul Wilkinson

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The Node is back on the road in 2014, and our first stop this year is the city of dreaming spires- Oxford in the UK!
 
 
Cat, the Node community manager, will be giving two talks on Monday the 13th of January:

‘How to use social media to promote and communicate your science’
2pm, Sherrington Library, Department of Physiology, Anatomy and Genetics (University of Oxford)

‘A career in publishing and science communication’
5pm, EPA seminar room, Sir William Dunn School of Pathology (University of Oxford)

Cat will also participate in the Research Career Pathways event at Oxford Brookes University on Tuesday afternoon.

Are you based in Oxford? Cat would love to meet you and hear your thoughts on the Node! Drop us an email if you would like to meet for a chat, or simply come to one of the talks! Looking forward to meet you!

(1 votes)

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CALL FOR A GROUP LEADER POSITION AT THE DEVELOPMENTAL BIOLOGY INSTITUTE OF MARSEILLE

The Developmental Biology Institute of Marseille (IBDM) is a leading research institute in Europe, with a unique focus on the study of developmental systems with interdisciplinary approaches using a wide range of animal models (Drosophila, Xenopus, C. elegans, chick, rat and mouse).

The IBDM is composed of 18 research groups and 5 scientific core facilities and benefits from the rich scientific environment of the Luminy campus of Aix-Marseille University. The overall research activity developed at the IBDM is at the crossroads of different fields: cell biology, development, evolution, neurobiology, physiology and biophysics. The connections and complementarity between these themes result in a strong scientific coherence of the overall research developed in the IBDM.

The teams employ transversal approaches and complementary strategies to understand how the instructions encoded in the genome are interpreted and translated to build structures (cells, tissues, organs) that perform specific functions, how these processes are regulated and integrated at the level of the whole organism and how their deregulation can lead to pathologies. A priority is to favor interdisciplinarity through the integration of new and original approaches that create conceptual and technical interfaces.

Please visit our website for more information.

We are looking for outstanding candidates who will complement the existing strengths of the Institute and develop an innovative and internationally competitive research program. Scientific excellence will be given the highest priority in the selection of the successful candidate.

This is a non-teaching position and knowledge of French is not required. The candidate will support his or her research by extramural funding * such as ERC, ATIP-Avenir or ANR.

Applicants should send in a single pdf file, a curriculum vitae, a list of publications, a 2 page summary of research achievements and projects in English, and the names and contacts of three references to the IBDM Director André Le Bivic (andre.le-bivic@univamu.fr) before the 30th of January 2014.

IBDM: CNRS / AIX-MARSEILLE UNIVERSITY, FRANCE

* Note from the contributor: All of these funding mechanisms are extremely competitive and difficult to obtain. ATIP-Avenir is the combined name of nationally awarded startup packages for promising young independent researchers from the CNRS or INSERM, now joint for this purpose. It is considered prestigious within France, though the prestige wears off after a few years. But the application deadline for an award in the second half of 2014 was the end of November, 2013.

ERC funding can be sought with the assistance of offices both at Université Aix-Marseille and at the CNRS (starting grants – success rate around 9% – or for established scientists, once cited on average at 14%, but that seems high nowadays from my anecdotal experience).

The ANR is the French national funding agency and it does help to understand written French to apply, though it’s not strictly required. 2013’s calls are here: not many are applicable to developmental biology. Success rates are hard to come by but from the POV of a mid-career developmental biologist, it’s not impossible and not easy. I guesstimate around 10%, too.

There aren’t other large extramural grant programs for research in France, of which I’m aware, relevant to developmental biology. There is international funding such as the Human Frontier Science Program but only you can see if it’s relevant to your case, when applying to such an offer.

On the plus side, such a position at the IBDM has a high chance of translating ultimately into tenure as a French civil servant. On the down side, if you don’t arrive with your own operating budget, it will be difficult to get going again at the standard at which you were recruited initially from elsewhere.

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Happy new year everyone!

To wrap up 2013 we had a look at our stats to find what out which were the most popular posts of the last year. 2013 saw the usual varied mix of news, research, meeting and discussion posts, so there was a lot to read!

Most viewed posts:

1- There and back again– Kara’s account of returning to the bench after working as an editor

2- Overly honest methods– a collection of the best tweets with this popular hashtag

3- Where scientists fear to tread– Caroline’s account of how ‘alternative’ careers are perceived

4- Breakthrough Prize floors winners with sheer amount of money– Eva commented on this newly established prize

5- A website for Postdocs and PhDs– the PostPostDoc website

Best rated posts:

1- There and back again– not only the most viewed but also the best rated!

2- The end of Biology?– Thomas’ thought provoking piece discussing some of the issues of science

3- Cellular Reincarnation– A literary interpretation of cellular reprogramming

4- A day in the life of.. a zebrafish lab

5- An interview with Alejandro Sánchez Alvarado

Other highlights:

2013 was a year that saw many people writing about their research and discussing their recent papers. Some of the most popular research posts this year included Making sense of Wnt signaling and a post by the University of Chicago journal club on the limb-to-fin transition. As has been the case in the past, our image competitions, such as our stem cell image competition or those featuring images from the Woods Hole course, have been extremely popular.

This last year also saw the beginning of two new series on the Node. A day in the life provides an account of a typical day in developmental biology labs working on different model organisms, and we have already covered many of the classical model systems. Our outreach series has already provided many case studies of outreach, as well as activity suggestions that you can try in your own outreach projects. Both series are continuing in 2014, so keep an eye out for more posts! We also launched a photography competition as part of our current outreach series- do participate for a chance to win a £50 Amazon voucher!

The Node is your community blog, and could not exist without your participation. So a big thank you to all of you who wrote, commented, rated or simply read the Node posts in 2013. We look forward to another exciting year of developmental biology in 2014!

Image: Andrea Pavanello (wikimedia commons)

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Currently, more and more women delay having children because of pursuing higher educational and career aspirations, as well as changing cultural norms. Unfortunately their eggs become susceptible to chromosome mis-segregation as a consequence of maternal aging. This would generate aneuploid embryos, so causing increased and birth defects (Jones and Lane, 2013). However, the actual ways in which chromosome segregation errors occur remain elusive, due to a lack of direct observations of the events as they happen. Live-cell tracking of chromosomes would be the infertility most appropriate technique to answer these questions, however with only chromosomal histone labeling, previous studies failed to follow any detailed dynamics of individual bivalents (Chiang et al., 2010; Lister et al., 2010).

In our recently published paper in Development (Yun et al., 2014), we applied a chromosome-tracking approach to examine bivalent dynamics in oocytes of aged mice during the entire period of meiotic maturation, by labeling both the bivalents and their kinetochores. By tracking, we have managed to reduce the intensity of imaging to such an extent that we can follow the movements of individual bivalents with a temporal resolution of 2 minutes continuously over a 12-15 hour time window, without any noticeable loss in rates of meiotic maturation (Movie 1). In so doing we have been able to catalogue the movements of bivalents and kinetochores in a way not previously performed, and establish the effects of maternal aging on chromosome dynamics in the first meiotic division (MI), through to metaphase II arrest (metII). Real-time tracking of bivalents in aged oocytes would be informative in the following aspects: 1) to determine if the process of bivalent congression necessary for faithful segregation is affected by age; 2) to determine the origin of single chromatids, which are commonly observed on metII eggs.

Using measurement of bivalent non-alignment when its displacement was >4 mm from the spindle equator (Lane et al., 2012), congression of all bivalents was achieved at least 3 hours ahead of anaphase onset independent of age, suggesting no gross malfunctioning of bivalent congression with age. However, we did observe more frequent weakly-attached bivalents in live aged oocytes, which had no apparent histone signal between the two sister chromatid pairs. Intriguingly, these bivalents did not undergo premature separation, but instead remained associated together all through MI. Despite the above observations in MI, the main defect with age was premature separation of dyads during metII arrest. The event was captured during imaging and occurred around 2 hours after anaphase I, as the metII spindle was assembling (Movie 2). The newly formed single chromatids oscillated about the spindle equator, presumably because they have only a single kinetochore that fails to establish simultaneous attachment to both spindle poles.

In conclusion, these data show that although considerable cohesion loss occurs during MI, its consequences are observed during meiosis II, when centromeric cohesion is needed to maintain dyad integrity, consistent with human studies that have shown a prevalence of pre-division in eggs from older women (Kuliev et al., 2011). The present work highlights that biopsy of the first polar body alone, which would have been normal in most aged oocytes here, may not be an effective screening method for aneuploidy.

References

Chiang, T., Duncan, F. E., Schindler, K., Schultz, R. M. and Lampson, M. A. (2010). Evidence that weakened centromere cohesion is a leading cause of age-related aneuploidy in oocytes. Current biology : CB 20, 1522-1528.

Jones, K. T. and Lane, S. I. (2013). Molecular causes of aneuploidy in mammalian eggs. Development 140, 3719-3730.

Kuliev, A., Zlatopolsky, Z., Kirillova, I., Spivakova, J. and Cieslak Janzen, J. (2011). Meiosis errors in over 20,000 oocytes studied in the practice of preimplantation aneuploidy testing. Reproductive biomedicine online 22, 2-8.

Lane, S. I., Yun, Y. and Jones, K. T. (2012). Timing of anaphase-promoting complex activation in mouse oocytes is predicted by microtubule-kinetochore attachment but not by bivalent alignment or tension. Development 139, 1947-1955.

Lister, L. M., Kouznetsova, A., Hyslop, L. A., Kalleas, D., Pace, S. L., Barel, J. C., Nathan, A., Floros, V., Adelfalk, C., Watanabe, Y. et al. (2010). Age-related meiotic segregation errors in mammalian oocytes are preceded by depletion of cohesin and Sgo2. Current biology : CB 20, 1511-1521.

Yun, Y., Lane, S. I. and Jones, K. T. (2014). Premature dyad separation in meiosis II is the major segregation error with maternal age in mouse oocytes. Development 141, 199-208.

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This PhD is part of the NERC funded Doctoral Training Partnership ACCE (Adapting to the Challenges of a Changing Environment). This is a partnership between the Universities of Sheffield, Liverpool, York, and the Centre for Ecology and Hydrology.

A PhD position is available in the Fraser laboratory at the University of Sheffield, Department of Animal and Plant Sciences and with co-supervisor Nathan Jeffery, Department of Musculoskeletal Biology at the University of Liverpool. The project is also in collaboration with Zerina Johanson, Department of Palaeontology, Natural History Museum, London.

Summary: The teeth of fishes and the integrated jaw apparatus are examples of extreme evolutionary modification that have responded to functional and adaptive shifts within the wider community. This novel project aims to identify shifts in biomechanical pressures on adult jaw and tooth type that is linked to changes in the development of the feeding system. Our integrative project surrounds the core question of how development contributes to novel evolutionary changes in trophic adaptation. This project will link biomechanical adaptation of morphology to novel developmental modifications of the jaw apparatus in fishes to ask whether having a novel dentition (e.g. beak-like dentition in pufferfishes) offers an adaptive advantage compared to more standard yet highly efficient dentitions e.g. Piranha. This project will utilize advanced techniques, including biomechanical computer simulations of hard-tissues built from enhanced microCT data. We will use nano-indentation analyses to observe changes of material properties in comparative groups of fishes linked to re-specification of conserved developmental genes in species with novel tooth phenotypes. The candidate will utilise developmental techniques (gene expression and manipulation) to understand how the genetic basis of tooth and jaw development and continuous tooth regeneration impact the evolution and biomechanical function of fish feeding systems.

Please visit the Department of Animal and Plant Sciences, University of Sheffield ACCE DTP website below for details of application.
http://www.sheffield.ac.uk/aps/prospectivepg/graduate-opportunities/accestudentships

The closing date for applications is January 20th 2014. For informal inquiries direct emails to Dr. Gareth Fraser: g.fraser@sheffield.ac.uk Lead Supervisor: Dr. Gareth Fraser, Dept. Animal and Plant Sciences, University of Sheffield. Co-supervisor: Dr. Nathan Jeffery, Dept. Musculoskeletal Biology, University of Liverpool. Project collaborator: Dr. Zerina Johanson, Dept. Palaeontology, Natural History Museum.

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