Why some vertebrates like salamanders and zebrafish are able to regenerate complex tissues while humans cannot is a question that has fascinated biologists for centuries. Understanding how and why regeneration occurs in these animals can inspire novel treatment strategies for regenerative medicine. At the cellular level, the regeneration process is driven by dynamic activities of cell migration, cell proliferation, and cell assimilation between old and new tissues. All of these events must be orchestrated in a precise order and at appropriate locations along the proximal-distal axis in order to restore a flawless, complex tissue (e.g. limbs or fins) from an amputation stump. With current imaging tools and platforms, it remains challenging to capture these dynamic, intricate cell behaviors in regenerating tissues from live adult vertebrates.

In 2012, my colleague in Dr. Ken Poss’s laboratory at Duke University, Vikas Gupta, had just successfully applied the “Brainbow” technique to the zebrafish heart to study cell behaviors during heart development and regeneration (Gupta and Poss, 2012). Since its debut in 2007, this elegant, multicolor cell labeling technique was mostly used to untangle the neuronal circuits in the brain (Livet et al., 2007). Vikas’s study demonstrated that this technique can also be applied to cell types other than nerve cells. At the time, biology aside, I was amazed by the beauty of the images he captured and started to wonder how I can apply this technique to fins, the tissue I study. The idea was that by tagging cells with diverse colors using the Brainbow cassette, I would be able to retrospectively determine contributions of distinctly labeled cells and their progeny in regenerating tissues, a key mechanistic question in understanding appendage regeneration. In addition, because fins are external, flat, and optically translucent, I might be able to uncover novel cell dynamics during regeneration by following these bar-coded cells in live animals. To label most cell types in fin tissue, I naively employed the ubiquitin promoter to drive the expression of Cre recombinase, while using the beta-actin2 promoter to drive the Brainbow cassette. To impose precise temporal control of Cre activity, I constructed a dual-inducible system that combines both the Tet-on system and an inducible Cre (CreERT2) in the transgene. The activity of Cre recombinase would require exogenous addition of both Doxycycline and Tamoxifen, limiting the possibility of leaky recombination. Such transgene design appeared to work nicely in injected, mosaic embryos.

Several months later when I began to screen through transgenic founders, I was at first disappointed to find that leaky recombination still occurred in many lines, and the expression domain of the Brainbow cassette was quite variable. However, I also noticed that progeny from one particular founder consistently displayed an unexpected, dazzling pattern (Figure 1) that was restricted to the outermost layer of the skin. Amazed by diverse hues displayed in this stable transgenic line, I assessed color stability of these labeled, post-mitotic cells by time-lapse imaging. Much to my surprise, multicolor tagging on this population of epithelial cells was rather stable, making tracing these cells over long time periods possible. Ken and I began to see that this “skin-bow” line may serve as a tool to study cell dynamics during skin turnover and regeneration. With hopes of tracing hundreds of cells in a large field of view, we were very fortunate to team up with two terrific quantitative biologists: Stefano Di Talia, who at the time just had established his laboratory at Duke, and Alberto Puliafito, a postdoctoral scientist in Luca Primo’s group in Italy to tackle this challenge. Alberto developed customized algorithms to segment our images, quantify and transform diverse cell behaviors that we just had a glimpse into compelling numbers.

With the skinbow system, we showed that regeneration of skin can be dissected into the most basic building block (i.e. cells), and each cell can be accurately monitored at the population level as regeneration takes place (Chen et al., 2016). Our findings identified diverse cell behaviors in response to different injuries that we would not have anticipated or discovered in fixed samples (click on video link below). The skinbow system provides a quantitative readout for studying these cell behaviors and their underlying mechanisms, many of which may be perturbed in aging, infected, or malignant skin tissues. As a proof of concept, we demonstrated that skinbow can be coupled with other transgenic lines to study cell-cell interactions during epithelial regeneration, or be employed as a screening platform to uncover molecular influences on certain cell behaviors. Among many future directions, I and others in the field are positioned to apply similar approaches (i.e. combination of cell barcoding, live imaging, and quantitative analysis) to illuminate activities of basal epithelial cells, bone cells, and mesenchymal cells in regenerating zebrafish tissues. The skinbow system might well be the first step to establish a complete, three-dimensional map of cell dynamics during vertebrate appendage regeneration. We merely scratched the surface of the subject at this point (literally!). New transgenic strains and analysis tools need parallel development to quantify cell behaviors in their respective z-positions, including in deep tissues. Nevertheless, I expect that new Brainbow cassettes that were recently developed in Jean Livet’s group (Loulier et al., 2014) would allow more flexibility in tagging and tracing different cell types in vivo, as now one can choose to paint either entire cells, or just nuclei and/or cell membranes in multicolor.

One thing I have learned to appreciate from this project is to be always on the lookout for unexpected findings, which can turn out to be more colorful than your best-laid plans.

See videos at: https://www.youtube.com/watch?v=xCNz1OHQ30E

More images at: https://www.flickr.com/photos/nihgov/26064937482/in/album-72157659401055954/

NIH director’s blog: https://directorsblog.nih.gov/2016/03/31/snapshots-of-life-fish-awash-in-color/

Duke Today: https://today.duke.edu/2016/03/zebrafish

The Economist: http://www.economist.com/news/science-and-technology/21695380-epidermis-now-comes-technicolor-rainbows-beginning

References:

Chen, C.H., Puliafito, A., Cox, B.D., Primo, L., Fang, Y., Di Talia, S., and Poss, K.D. (2016). Multicolor Cell Barcoding Technology for Long-Term Surveillance of Epithelial Regeneration in Zebrafish. Dev Cell 36, 668-680.

Gupta, V., and Poss, K.D. (2012). Clonally dominant cardiomyocytes direct heart morphogenesis. Nature 484, 479-484.

Livet, J., Weissman, T.A., Kang, H., Draft, R.W., Lu, J., Bennis, R.A., Sanes, J.R., and Lichtman, J.W. (2007). Transgenic strategies for combinatorial expression of fluorescent proteins in the nervous system. Nature 450, 56-62.

Loulier, K., Barry, R., Mahou, P., Le Franc, Y., Supatto, W., Matho, K.S., Ieng, S., Fouquet, S., Dupin, E., Benosman, R., et al. (2014). Multiplex cell and lineage tracking with combinatorial labels. Neuron 81, 505-520.

(2 votes)

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Applications are invited for a 2 year Wellcome Trust funded Research Assistant position to join an international team in the Department of Genetics in central Cambridge. The project is led by Dr Ben Steventon and is aimed towards understanding the role of gene expression heterogeneity in the control of neural/mesodermal cell fate decisions during the elongation of the posterior body axis in zebrafish embryos. For further details on this position please look here.  For further details on our research, please visit steventonlab.wordpress.com

The post-holder will be involved the generation of zebrafish transgenic lines that will enable the imaging of gene-expression dynamics in vivo. In addition, they will utilize cutting-edge imaging techniques to quantify gene expression levels in situ.

The successful candiate will be a highly motivated and well-organised individual with a first degree in biological or biomedical sciences and experience in molecular biology. Experience in zebrafish genetics would be an advantage.

Fixed-term: The funds for this post are available for 2 years in the first instance.

The University values diversity and is committed to equality of opportunity.

The University has a responsibility to ensure that all employees are eligible to live and work in the UK.

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The Evolutionary and Functional Genomics Lab led by Josefa González is seeking a highly motivated postdoctoral researcher to join our research team at the Institute of Evolutionary Biology (CSIC-UPF).
The postdoctoral researcher will work on a project funded by a European Research Council Consolidator Grant that aims at identifying the genetic basis, the molecular mechanisms, and the functional traits relevant for environmental adaptation.
The postdoctoral researcher will be responsible for the in silico characterization of candidate adaptive mutations identified in natural populations of Drosophila melanogaster. Among others, the tasks involved in the postdoctoral research project will be to identify pathways under selection, and to analyze the expression and the epigenetic changes of genes nearby the candidate adaptive insertions.
A PhD in Populations Genetics or a related field, good programming skills, and good writing skills are required. Previous postdoctoral experience will be considered.
We offer a full-time position for 2 years with the possibility of extension. Salary will depend on the experience of the candidate.
Starting date September 2016 but alternative dates can be discussed.

Application
Please send your CV and a brief letter of motivation before the 5th May 2016 to: josefa.gonzalez@ibe.upf-csic.es

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It was on the 7^th of February of 2016 when 20 leading scientists from all over the world headed to the historic Wiston House in West Sussex, England, to spend four days in focused atmosphere discussing new insights in cardiovascular research: the workshop for Transdifferentiation and Tissue Plasticity in Cardiovascular Rejuvenation. Supported by the Company of Biologists, Brian Black and Jim Martin brought together experts of the field of heart development, regeneration and tissue engineering with the aim to discuss new approaches and recent findings to improve cardiac repair. In addition to the 20 senior investigators, also 10 early-stage scientists (PhD students, postdocs, and junior PIs) were selected to participate. For me in particular, with my PhD recently completed, joining this event was a great honour and a unique opportunity. I excitedly anticipated the four intense workshop days with great interest.

With the heart being such a complex and specialized tissue, repair of cardiac muscle in human is difficult to achieve. In the workshop, we were introduced to and discussed a wide range of ongoing efforts, including genomic regulation of cardiomyocyte differentiation, environmental factors involved in cardiac regeneration, as well as the contribution of different cell types in this process. Although I cannot allude to all the talks here, all participants agreed about the outstanding scientific quality of the work presented in this meeting.

Liz Robertson opened the first session of the workshop on Sunday afternoon recapitulating the origin of cardiac progenitors during embryogenesis and explaining the involvement of Eomesodermin in cardiac mesoderm specification in the mouse embryo (Costello et al., 2011). To understand myocardial differentiation, it is fundamental to define transcription factors and epigenetic modifications that specify cardiac lineages. Benoit Bruneau introduced his lab’s latest study on the coordination of three cardiac transcription factors (Nkx2.5, Tbx5 and Gata4) in the regulation of cardiac gene expression and differentiation (Luna-Zurita et al., 2016). Laurent Dupays described the interaction of the two transcription factors Meis and Nkx2.5 on a specific enhancer sequence (Dupays et al., 2015). Moreover, Brian Black explained new findings from his group about the Mef2c transcriptional regulation machinery in cardiomyocytes.

We also learned about specific approaches to elucidate the functions of distinct cellular factors active in cardiomyocytes. Guo Huang is currently investigating fetal cardiac genes, which reactivate cell cycle re-entry of adult heart muscle cells for potential regenerative repair after myocardial infarction. Jim Martin reported the implication of the Hippo pathway in cytoskeletal remodelling of cardiomyocytes in the injured heart (Morikawa et al., 2015). Finally, Kathy Ivey introduced how to study human iPSC-derived cardiomyocytes to better understand protein signalling and interaction networks as well DNA-occupancy in cellular differentiation.

Another focus during this workshop was the understanding of environmental factors, which impact cardiomyocyte behaviour and fate during cardiac repair. We learned from Eldad Tzahor how the stiffness of the extracellular matrix affects the differentiation state of cardiomyocytes (Yahalom-Ronen et al., 2015). Ahmed Mahmoud described the implication of cardiac innervation and Neuregulin signalling in the regulation of cardiomyocyte proliferation in the regenerating neonatal mouse and zebrafish heart (Mahmoud et al., 2015). The fact that many different cell types are crucial for cardiac regeneration was demonstrated by Paul Riley and Nadia Rosenthal. Nadia nicely illustrated the cellular composition of the heart and discussed recent work that demonstrates the fundamental role of macrophages during cardiac repair and regeneration (Pinto et al., 2016), while Paul Riley explained the different origins and the development of lymphatic vessels in the heart and described how this developmental program is reactivated after myocardial infarction (Klotz et al., 2015). Another approach was described by Enzo Porrello, who is seeking to understand the differences between cardiac cells at different stages of life, to unveil the mechanisms that impede adult cardiac regeneration.

Multiple talks presented studies using the zebrafish, an important model of cardiac development and regeneration due to its remarkable regenerative capacity and its transparency during embryogenesis. Didier Stainier illustrated how cardiomyocytes delaminate from the compact layer to form trabeculated myocardium in the zebrafish embryo (Staudt et al., 2014). Karina Yaniv showed beautiful movies displaying the origin of lymphatic vessels during development (Nicenboim et al., 2015). Christian Mosimann presented live imaging that traces back the cardiovascular lineages to the lateral plate mesoderm (Mosimann et al., 2015) and explained what we could learn about cardiac diseases by modelling human patient mutations in zebrafish. Ken Poss, whose lab is interested in the mechanisms of heart and fin regeneration, guided us through his journey in searching for regenerative cellular programs in the zebrafish (Kang et al., 2016). Further, Nadia Mercader spoke about distinct populations of cardiomyocytes in the early and the adult zebrafish heart and injury studies to decipher the mechanisms of myocardial regeneration.

Another key topic discussed in this workshop was how we can investigate the underlying causes of human cardiac diseases in more depth. Alessandra Moretti showed one example of how her lab studies the cause of Arrhythmogenic right ventricular dysplasia using patient-derived iPSCs. Deepak Srivastava further reported new findings about the molecular consequences of human GATA4 mutations obtained by studying iPSCs-derived cardiomyocytes from patients. An impressive finding was reported by Eric Olson: he explained how his lab achieved the repair of mutations in the Dystrophin gene, the cause of Duchenne muscular dystrophy, by CRISPR/Cas 9 technology in mice in vivo (Long et al., 2016).

I was personally very intrigued to hear about the work of the bio-engineers participating in this workshop. Nenac Bursac illustrated how his lab gains insights into cardiomyocyte functions by using in vivo assays of cardiomyocyte patches. Moreover, from Molly Stevens we learned about the versatile use of nano needles, which can deliver substances to cells or even make measurements (Chiappini et al., 2015).

In my opinion, and I am sure all participants would agree, this workshop was a tremendous success. Fascinating data, of high scientific value were presented and openly discussed. For me, this workshop was a unique experience; I met experts in the field of cardiovascular research and learned about a vast range of experimental approaches. Moreover I had the opportunity to present and discuss our latest results on the endocardial dynamics in zebrafish heart regeneration. Finally the guided tour through the amazing 16^th century historical Wiston House, the windy walk in the beautiful Sussex countryside, and the experience of watching the Super Bowl for the first time, completed the great experience of the Transdifferentiation and Tissue Plasticity in Cardiovascular Rejuvenation workshop.

Here is a short video put together by the Company of Biologists on this workshop:

I am grateful to Christian Mosimann for comments on the text.

References

Chiappini, C., De Rosa, E., Martinez, J. O., Liu, X., Steele, J., Stevens, M. M. and Tasciotti, E. (2015). Biodegradable silicon nanoneedles delivering nucleic acids intracellularly induce localized in vivo neovascularization. Nature materials 14, 532-539.

Costello, I., Pimeisl, I. M., Drager, S., Bikoff, E. K., Robertson, E. J. and Arnold, S. J. (2011). The T-box transcription factor Eomesodermin acts upstream of Mesp1 to specify cardiac mesoderm during mouse gastrulation. Nature cell biology 13, 1084-1091.

Dupays, L., Shang, C., Wilson, R., Kotecha, S., Wood, S., Towers, N. and Mohun, T. (2015). Sequential Binding of MEIS1 and NKX2-5 on the Popdc2 Gene: A Mechanism for Spatiotemporal Regulation of Enhancers during Cardiogenesis. Cell reports 13, 183-195.

Kang, J., Hu, J., Karra, R., Dickson, A. L., Tornini, V. A., Nachtrab, G., Gemberling, M., Goldman, J. A., Black, B. L. and Poss, K. D. (2016). Modulation of tissue repair by regeneration enhancer elements. Nature.

Klotz, L., Norman, S., Vieira, J. M., Masters, M., Rohling, M., Dube, K. N., Bollini, S., Matsuzaki, F., Carr, C. A. and Riley, P. R. (2015). Cardiac lymphatics are heterogeneous in origin and respond to injury. Nature 522, 62-67.

Long, C., Amoasii, L., Mireault, A. A., McAnally, J. R., Li, H., Sanchez-Ortiz, E., Bhattacharyya, S., Shelton, J. M., Bassel-Duby, R. and Olson, E. N. (2016). Postnatal genome editing partially restores dystrophin expression in a mouse model of muscular dystrophy. Science 351, 400-403.

Luna-Zurita, L., Stirnimann, C. U., Glatt, S., Kaynak, B. L., Thomas, S., Baudin, F., Samee, M. A., He, D., Small, E. M., Mileikovsky, M. et al. (2016). Complex Interdependence Regulates Heterotypic Transcription Factor Distribution and Coordinates Cardiogenesis. Cell 164, 999-1014.

Mahmoud, A. I., O’Meara, C. C., Gemberling, M., Zhao, L., Bryant, D. M., Zheng, R., Gannon, J. B., Cai, L., Choi, W. Y., Egnaczyk, G. F. et al. (2015). Nerves Regulate Cardiomyocyte Proliferation and Heart Regeneration. Developmental cell 34, 387-399.

Morikawa, Y., Zhang, M., Heallen, T., Leach, J., Tao, G., Xiao, Y., Bai, Y., Li, W., Willerson, J. T. and Martin, J. F. (2015). Actin cytoskeletal remodeling with protrusion formation is essential for heart regeneration in Hippo-deficient mice. Science signaling 8, ra41.

Mosimann, C., Panakova, D., Werdich, A. A., Musso, G., Burger, A., Lawson, K. L., Carr, L. A., Nevis, K. R., Sabeh, M. K., Zhou, Y. et al. (2015). Chamber identity programs drive early functional partitioning of the heart. Nature communications 6, 8146.

Nicenboim, J., Malkinson, G., Lupo, T., Asaf, L., Sela, Y., Mayseless, O., Gibbs-Bar, L., Senderovich, N., Hashimshony, T., Shin, M. et al. (2015). Lymphatic vessels arise from specialized angioblasts within a venous niche. Nature 522, 56-61.

Pinto, A. R., Ilinykh, A., Ivey, M. J., Kuwabara, J. T., D’Antoni, M. L., Debuque, R., Chandran, A., Wang, L., Arora, K., Rosenthal, N. A. et al. (2016). Revisiting Cardiac Cellular Composition. Circulation research 118, 400-409.

Staudt, D. W., Liu, J., Thorn, K. S., Stuurman, N., Liebling, M. and Stainier, D. Y. (2014). High-resolution imaging of cardiomyocyte behavior reveals two distinct steps in ventricular trabeculation. Development 141, 585-593.

Yahalom-Ronen, Y., Rajchman, D., Sarig, R., Geiger, B. and Tzahor, E. (2015). Reduced matrix rigidity promotes neonatal cardiomyocyte dedifferentiation, proliferation and clonal expansion. eLife 4.

(3 votes)

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Every year, the British Society for Developmental Biology (BSDB) awards the Beddington Medal to the best PhD thesis in developmental biology. The 2016 award went to Elena Scarpa, who did her PhD with Roberto Mayor at University College London (UCL). We caught up with Elena at the BSCB/BSDB Spring meeting, and we asked her about her thesis work on the neural crest and what she is doing now.

Congratulations on winning the Beddington Medal. What does this prize mean to you?

I am very happy about this prize. I moved from Italy to the UK to do my PhD, which was a big change for me. I knew my science background was good, but at UCL I was surrounded by people who had come from excellent universities, like Cambridge or UCL, and my university in Italy was not recognised in the same way. Some of the other students in my PhD course were foreigners as well, but they had studied in the UK. So receiving this medal is a big achievement.

This prize also recognises the communal effort that went into this project. It was a lot of work, not just from me but also from the people I collaborated with and helped me. I think this prize also reflects well on Roberto. I am the second student from his lab that wins it, showing that his students are doing well and that he is a good supervisor.

Can you tell us a bit more about the lab where you did your PhD?

I did my PhD in Roberto Mayor’s lab at UCL. The lab works on neural crest, mostly in Xenopus but also in zebrafish. The lab started out by working on induction and specification of the neural crest, but in the last few years the majority of the lab has worked on different aspects of neural crest migration. The lab had a very nice environment. There were many students, so it was a lot of fun.

It seems that contact inhibition is on a winning streak, since last year’s winner of the Beddington Medal, John Robert Davis, was also working on contact inhibition. What is contact inhibition and in what contexts is it important?

Contact inhibition is the process by which two migrating cells interact and change their direction of motion after contact. This phenomenon was discovered by Abercrombie at UCL. Roberto likes to say that Abercrombie was based in the same room as the Mayor lab, but I am not sure this is true! Abercrombie and his colleague Heaysman observed that contact inhibition was related with the ability of malignant cells to invade other tissues. Their work was very nice but descriptive, and contact inhibition and its molecular mechanisms did not receive much attention for several years.

In 2008 Roberto’s lab published a paper about contact inhibition of locomotion, showing that this process is required for neural crest directionality, and hinting at the molecular mechanism behind it. It showed that cell-cell contacts, planar cell polarity and cadherins are required. However, what I think is interesting about contact inhibition is that it mediates many different types of cell behaviour during development. In the neural crest it mediates collective behaviour (in combination with other processes). However, as John showed in hemocytes, and also in neurons, it can mediate dispersion. In addition, it can be used by certain cancer cells as a driving force for invasion. So the presence or lack of contact inhibition can really change how cells interact with each other.

What was your thesis project about, and what were your major findings?

Contact inhibition is the process by which cells separate, but many other cells types, such as epithelial cells, stay together and make stable contacts. We wanted to understand better the nature of mesenchymal cell-cell interactions. Are mesenchymal cell-cell interactions like contact inhibition intrinsically different because cells are unable to form a junction, or is there something in the junction itself that changes the behaviour of the cells?

My main finding was that, at least in the neural crest, cells that undergo contact inhibition do not express different cadherins. The ability of cadherins to recruit the cadherin complex or the actin complex is very similar, but there is a difference in their ability to polarise the activity of small GTPases, polarising the motility of the cells. This adds to what was already previously known. In the neural crest and in cancer it is well known that E-cadherin is down regulated, and this seems to correlate with the ability of the cells to invade. However, the idea that loss of contact allows invasion does not really make sense. Cells that are unable to make contacts with other cells cannot interact properly with their environment, so in vivo this does not favour migration. Our paper showed that there are other changes in the way cells form protrusions and interact with their environment that lead to scattering and active migration, rather than just the cell-cell contact itself. Other papers had proposed this idea before, but we were able to put it in a developmental framework.

You mentioned collaborators. Which labs did you collaborate with?

Within the lab I originally worked with Eric Theveneau, and also with András Szabó who is a modeller and helped me with the quantitative analysis of traction forces.

During my project we used FRET to obtain more information about the dynamics of how contact inhibition is regulated. For this I collaborated with Maddy Parsons at King’s College London, who had already collaborated with Roberto on other contact inhibition papers. I worked with Maddy a lot, and it took a lot of effort in the first two years of my PhD to get the FRET to work. It is very nice to collaborate with Maddy because she is very available, and happy to try different things.

I also established a collaboration for the optogenetics part of the project, since were not able to get it to work at UCL. During my PhD Eric moved to Toulouse, where Xiabo Wang , who previously adapted photoactivation in vivo during his postdoc in Denise Montell’s lab and used a photo-activatable form of Rac. He was familiar with the imaging conditions, so I did my experiments in their microscope in Toulouse. This was a great opportunity to learn a new technique, and I am going to develop optogenetics further in my postdoc. So this collaboration also helped me to define my interests.

You have moved from London to Cambridge for your postdoc. What are you working on now and how did you choose your new lab?

I now work in the lab of Benedicte Sanson at the department of Physiology, Development and Neuroscience. The lab studies the Drosophila embryo at the global level, examining how extrinsic forces generate collective rearrangements in the tissue. They focus mostly on germ band extension and parasegment boundary formation. My project is concerned with oriented cell division in the early Drosophila embryo. I am looking at how mechanical cues influence the orientation of cell division.

During my PhD I looked at cell migration, and how the traction on the substrate relates with the tension mediated by cell-cell interactions. For my postdoc I wanted to develop more my knowledge of tissue mechanics. The Sanson lab is a really good place to develop this interest because the Drosophila embryo is a really powerful system. It is very simple and very easy to access. They also have a great ongoing collaboration with Guy Blanchard from Richard Adam’s lab. Their tracking software is very powerful and allows very fine image analysis. You can track mesenchymal cells like the ones I studied during my PhD, but it is never very refined because it is very difficult to segment the cells. In epithelial cells this type of analysis is possible.

Did you find it difficult to change model organism?

It is different. In the beginning working with Drosophila didn’t make sense to me. I am used to Xenopus and microinjections. When you come into the lab on a Monday you don’t know exactly what will you do, and choose your experiment based on the quality of the embryos. Because you just microinject it isn’t necessary to plan ahead, but on the other hand you can’t do clean genetics. In Drosophila you have very powerful genetic tools which are very useful. However, you need to plan your experiments. You need to cross the flies, wait, then select them… It just didn’t make sense to me why I had to wait 2 weeks to see the membrane! But you get used to it.

Do you have any advice for new students?

Choose something that you really like and that really motivates you. You are going to spend a lot of time in the lab trying to solve problems, so you need a good question that you are passionate about. Also try to choose a good environment, where you will be supported and have interesting discussions. Finally, be creative. Sometimes you get stuck, and if you tried everything and it doesn’t work then you need to think outside the box. Be a bit brave and take some risks.

(4 votes)

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Location: University of Exeter

Research Group: James Wakefield (www.thewakefieldlab.com)

Salary scale: £19,828 – £25,023

Duration: 1st June 2016 – 31st August 2018 (with possible 3 yr renewal)

We wish to recruit a Research Technician to support the work of Dr James Wakefield.  The successful applicant will assist Dr Wakefield in the day-to-day management of the lab and work on an interdisciplinary project aimed at understanding the fundamental process of mitotic spindle formation in the model organism, Drosophila melanogaster. As such, this position represents an opportunity to work in a creative, collegiate and interdisciplinary research environment, making an essential contribution to an internationally-leading research programme.

The successful applicant will support a wide range of research activities, including supporting and training undergraduate and post-graduate research students, working closely with the PI to organise and co-ordinate aspects of lab-management, overseeing the culture and maintenance of Drosophilalaboratory stocks and undertaking their own independent research project. They will be enthusiastic, highly motivated and possess excellent verbal and written communication skills.

Applicants will possess a relevant first degree (BSc Honours) in Biological Sciences, Biochemistry, Genetics or a related subject and demonstrate sufficient knowledge of research methods and techniques to work within the established research programme. Applicants will be able to demonstrate skills in genetics, biochemical techniques, microscopy and cell biology. Previous experience working with Drosophila would be a distinct advantage.

For further information, please see

http://www.jobs.ac.uk/job/ANM107/research-technician/

Informal contact is encouraged:

email J.G.Wakefield@ex.ac.uk, or telephone (07966 660604)

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A postdoctoral position is available to study the developmental mechanisms that pattern differentiation during organ development.

Epithelial tubes often have a functional polarity written along their P-D axis, with specialised segments carrying out distinct physiological activities. With a handful of notable exceptions, we know little about how P-D axes and segment-specific differentiation are regulated during organogenesis.

We aim to understand the molecular and cellular mechanisms that pattern and maintain functional polarity along the P-D axis in a structurally simple, but functionally sophisticated epithelial tube: the Drosophila renal tubule.

Highly motivated applicants with a PhD and strong background in cell/developmental biology are encouraged to apply.

For informal inquiries about the position please contact Barry Denholm directly: Barry.Denholm@ed.ac.uk

Vacancy ref: 036068

https://www.vacancies.ed.ac.uk/

Closing date: 20-May-2016

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This cartoon was first published in the Journal of Cell Science. Read other articles and cartoons of Mole & Friends here.

Part I- ‘The imposter’

Part II- ‘The teaching monster’

Part III- ‘The Pact’

Part IV- ‘The fit’

Part V- ‘The plan’

Part VI- ‘FCTWAWKI’

Part VII- ‘Beaten and bruised’

Part VIII- ‘Money Matters’

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The origin of paired fins is a major unresolved issue in vertebrate evolutionary biology, and has been a topic of debate among palaeontologists, comparative anatomists and developmental biologists for over a century. Central to any question of “evolutionary origins” is the concept of homology: the sharing of features due to common ancestry. Homology may explain the existence of shared features between organisms (historical homology – e.g. the arms of a chimpanzee and the arms of a human are homologous because these structures have been retained from a common ancestor that possessed arms), or the existence of shared features within an organism (serial homology – e.g. the vertebral elements within a human, which exhibit a range of morphologies, but which nevertheless share a common underlying ground plan) (Roth, 1984; Wagner, 1989). In either case, homology reflects a continuity of anatomical, cellular, or genetic information, and provides a useful conceptual framework for investigating the evolutionary relationship among body plan features in distantly related taxa (Van Valen, 1982).

Cartilaginous fishes (sharks, skates, rays and holocephalans) are unique among living jawed vertebrates, in that they possess a series of skeletal appendages called branchial rays that project laterally from their gill arches (Fig. 1a). These branchial rays articulate with the gill arch cartilages in a way that is broadly reminiscent of the articulation between pectoral fins or limbs and the shoulder girdle, and this similarity led the comparative anatomist Carl Gegenbaur to propose that paired fins and limbs evolved by transformation of a gill arch (Gegenbaur, 1878) (Fig. 1b, c) – a hypothesis of serial homology that remains controversial to this day. Unfortunately, the fossil record currently tells us relatively little about the stepwise acquisition of paired fins during vertebrate evolution, so we decided to address this question from a developmental perspective. We were interested in determining whether the anatomical parallels that Gegenbaur noted between the gill arches of cartilaginous fishes and fins/limbs may reflect common underlying molecular patterning mechanisms in these organ systems. To this end, we conducted a series of experiments to investigate branchial ray patterning in embryos of an oviparous (egg-laying) cartilaginous fish, the little skate, Leucoraja erinacea (see video below for an overview of skate embryonic development).

It might be helpful to first briefly introduce the gill arch, and how the gill arches of cartilaginous fishes are unique relative to those of other jawed vertebrates. All vertebrate embryos possess a bilateral series of pharyngeal arches on the sides of their developing head (Fig. 2a), and the mesenchyme within these arches gives rise to much of the craniofacial skeleton. The first pharyngeal arch is called the mandibular arch, the second is called the hyoid arch, and this is followed by a variable number of gill arches (most cartilaginous fishes have five gill arches). Primitively, the mandibular arch gave rise to the jaw skeleton, the hyoid arch gave rise to the skeletal apparatus that suspends the jaw from the braincase, and the gill arches give rise to the skeletal supports of the respiratory apparatus of the gills. These skeletal derivatives are conserved in a relatively primitive organisation in cartilaginous fishes, but are also present in other vertebrates (though often in a derived state – for example, the skeletal derivatives of the “gill arches” of mammals give rise to the laryngeal skeleton). Uniquely, though, in cartilaginous fishes, once the hyoid and gill arches have formed, they undergo a lateral expansion (Fig. 2b), and give rise to an additional set of skeletal elements – the branchial rays – that project laterally from the arches (Gillis et al., 2009). These branchial rays ultimately provide skeletal support to the fleshy flaps that protect the gills of cartilaginous fishes. As appendages, branchial rays must be patterned along the proximodistal axis (as they expand laterally) and also along the anterior-posterior axis (branchial rays exhibit a pronounced anterior-posterior polarity, and articulate proximally along the posterior margin of the hyoid and gill arch cartilages). In our paper (Gillis and Hall, 2016), we were interested in testing whether skate gill arches deploy similar axial patterning mechanisms as do fins and limbs (as would be predicted by an hypothesis of gill arch-fin/limb serial homology).

In this study, we focused our attention on the sonic hedgehog (Shh) signaling pathway, as the axial patterning function of this pathway during limb development is very well understood. Decades of experimental embryological and molecular investigation have revealed that Shh signaling is important in both anterior-posterior patterning of the limb, and in the proliferative expansion of limb skeletal progenitors. Classical chick embryo experiments by Saunders and Gasseling (1968) demonstrated that upon transplantation of donor posterior limb bud mesenchyme to the anterior region of a host limb bud, the resulting limb will form an additional set of digits that are oriented mirror-image to the normal digits (Fig. 3a). This posterior limb bud signaling centre is known as the zone of polarizing activity (ZPA). Twenty five years later, the Tabin lab demonstrated that Shh was the polarizing signal that was being secreted by the ZPA (Fig. 3b,c), and that misexpression of Shh in the anterior limb bud was sufficient to induce the ectopic, mirror-image digits noted by Saunders and Gasseling (Fib. 3d,e) (Riddle et al., 1993). In addition to this anterior-posterior patterning role, Shh signaling is also required for the expansion of limb endoskeletal progenitors, and progressively earlier loss of Shh signaling during limb development results in a progressively more profound deletions of distal limb skeletal elements (as shown, for example, by the genetic deletion of Shh from mouse limb buds, or by pharmacological manipulation of hedgehog signaling in chick and salamander) (Fig. 3f) (Towers et al., 2008; Zhu et al., 2008; Stopper and Wagner, 2007). Does Shh signaling function in a similar manner during the development of skate branchial rays?

We first sought to determine whether Shh signaling components were expressed during skate gill arch development. mRNA in situ hybridization experiments revealed that, indeed, Shh is expressed in a polarized pattern during the development of the skate hyoid and gill arches – initially in the posterior epithelium of each arch, and eventually in an epithelial stripe along the leading edge of the hyoid and gill arches as they undergo lateral expansion (Fig. 4). When we looked at the expression of Ptc2 (a transcriptional readout of Shh signaling), to determine which tissues are responding to this Shh signal, we observed expression in both the distal gill arch epithelium, and in the mesenchyme beneath the Shh expression domain (Fig. 4). One key difference, of course, is that in skate gill arches, Shh is expressed in posterior-distal epithelium, while in the limb bud, Shh is expressed in posterior mesenchyme. However, in both cases, Shh signal is transduced in overlying epithelium and distal mesenchyme (so although the source of the signal is different between these appendages, the responding tissues are similar).

In order to determine whether Ptc2+ mesenchymal cells (i.e. mesenchymal cells responding to Shh signal) ultimately contribute to the branchial ray skeleton, we conducted a fate mapping experiment. Over the past several years, I have developed protocols for the experimental manipulation and long term in/ex ovo culture of skate embryos, and this now allows us to conduct targeted embryonic manipulations (e.g. by microinjecting and surgically manipulating specific regions of the embryo) and to focally label populations of embryonic cells, in order to trace the long-term fates of their progeny (Fig. 5a,b). For this experiments, we microinjected the lipophilic dye CM-DiI immediately subjacent to the Shh-expressing epithelium of the gill arch, so that we could assess the contribution of these cells (and their progeny) to the gill arch skeleton. CM-DiI is readily incorporated into cell membranes, and is retained in daughter cells through mitosis (although diluted somewhat with each round of cell division). Importantly, though, CM-DiI will persist through fixation and paraffin sectioning, and so upon skeletal differentiation, we can use fluorescent microscopy on thin sections to recover even very small specks of membrane-localized CM-DiI (indicating decent from mesenchymal cells that were Shh-responsive earlier in development). These experiments demonstrated that Shh-responsive gill arch mesenchyme does contribute to branchial rays (Fig. 5c), and suggest that this signaling pathway may be directly influencing the behaviour and fate of branchial ray progenitors.

Finally, to test the function of Shh signaling during skate branchial ray development, we conducted a series of in ovo pharmacological treatments. Skate embryos develop in large, leathery egg shells, and are amenable to bath treatment by in ovo injection of small molecules. We used cyclopamine – a small molecular inhibitor of the hedgehog signaling pathway – to inhibit Shh signaling at different stages of gill arch development, and to test for stage-specific roles for Shh signaling in bronchial ray patterning. We chose three stages for treatment: stage 22 (gill arches have still formed, and Shh is expressed in posterior arch epithelium), stage 27 (gill arches are undergoing lateral expansion, with Shh signaling resolved to an epithelial stripe along the leading edge of the expanding arch) and stage 29 (just prior to the condensation of the gill arch endoskeleton). Interestingly, upon manipulation of Shh signaling at these different stages of gill arch development, we observed branchial ray defects that were broadly reminiscent of the skeletal defects observed upon manipulation of Shh signaling during limb development. For example, we observed that progressively earlier inhibition of Shh signaling resulted in a progressively greater deletion of branchial rays (i.e. cyclopamine treatments at stages 22 and 27 resulted in a significant reduction in the number of branchial rays on each arch, while treatment at stage 29 resulted in no significant difference in branchial ray number) (Fig. 6a). We also observed that cyclopamine treatment at stage 22 resulted in loss of anterior-posterior axis specification (i.e. the few branchial rays that did form articulate down the midline of the arch, rather than along the posterior margin of the arch) while cyclopamine treatment at stages 27 or 29 had no effect on the anterior-posterior axis (Fig. 6b). It therefore appears that, as in the limb bud, Shh signaling functions initially in skate gill arches to establish the anterior-posterior axis, and subsequently to maintain proliferative expansion of branchial ray endoskeletal progenitors.

So what does this shared role for Shh signaling in gill arch and limb bud patterning mean? It is possible that limbs share a patterning mechanism with gill arches because these structures are, indeed, transformational homologues (i.e. fins and limbs evolved by transformation of a gill arch in an ancestral vertebrate, as proposed by Gegenbaur). However, it may also be that gill arches and fins/limbs have independently recruited a deeply conserved “core” appendage patterning mechanism (i.e. parallel evolution, leading to serial homology), or that gill arches and fins/limbs are convergently using the Shh signaling pathway for similar purposes. Only palaeontological data can tell us about anatomical transitions, and such data are needed in order to formally test Geganbaur’s hypothesis of gill arch-paired fin transformational homology. However, it is now clear that some of the anatomical parallels that led Gegenabur to propose his gill arch hypothesis of fin origins reflect common underlying patterning mechanisms, and further investigation of the molecular basis of branchial ray patterning in cartilaginous fishes will allow us to determine whether these common mechanisms are the result of parallel evolution or convergence. I think that this study sets out an exciting path forward to address the origin and evolution of paired appendages in vertebrates, and highlights how complementary palaeontological and developmental approaches are needed in order to truly address the big, unanswered questions in vertebrate body plan evolution.

References

Gegenbaur, C (1878) Elements of Comparative Anatomy. London, UK: Macmillan.

Gillis, J.A., Dahn, R.D. and Shubin, N.H. (2009) Chondrogenesis and homology of the visceral skeleton in the little skate, Leucoraja erinacea (Chondrichthyes: Batoidea). J. Morphol. 270, 628-643.

Gillis, J.A. and Hall, B.K. (2016) A shared role for sonic hedgehog signalling in patterning chondrichthyan gill arch appendages and tetrapod limbs. Development 143, 1313-1317.

Owen, R. (1866) Anatomy of Vertebrates I. Fishes and Reptiles. London, U.K.: Longmans, Green, and Co.

Riddle, R.D., Johnson, R.L., Laufer, E. and Tabin, C.J. (1993) Sonic hedgehog mediates the polarizing activity of the ZPA. Cell 75, 1401-1416.

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Stopper, G.F. and Wagner, G.P. (2007) Inhibition of Sonic hedgehog signaling leads to posterior digit loss in Ambystoma mexicanum: parallels to natural digit reduction in urodeles. Dev. Dyn. 236, 321-331.

Towers, M., Mahood, R. Yin, Y. and Tickle, C. (2008) Integration of growth and specification in chick wing digit patterning. Nature 452, 882-886.

Van Valen, L.M. (1982) Homology and causes. J. Morphol. 173, 305-312.

Wagner, G.P. (1989) The biological homology concept. Annu. Rev. Ecol. Syst. 20, 51-69.

Zhu, J., Nakamura, E., Nguyen, M.T., Bao, X., Akiyama, H. and Mackem, S. (2008) Uncoupling sonic hedgehog control of pattern and expansion of the developing limb bud. Dev. Cell. 14, 624-632.

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