The neural crest arises between neural and non-neural ectoderm and represents a somatic cell type with unique properties of multipotency. The neural crest cells (NCCs) migrate throughout the body and differentiate into a diverse array of cell types and tissues including the peripheral nervous system, enteric and sensory nervous system, Schwann cells, skin melanocytes, as well as connective tissues. The neural crest has major clinical relevance since it is disproportionately involved in both inherited and acquired developmental abnormalities termed neurocristopathies.

NCCs migration, development, and differentiation into various tissues have been studied in vivo in avian embryos, in studies pioneered by Nicole Le Douarin. Donor quail neural crest tissues were grafted into similar regions of stage-matched developing chick embryos, to generate quail-chick neural crest chimeras in ovo. The cells of the two species are easy to distinguish based on the ability to identify interphase nucleus of the quail, in which a large amount of heterochromatic DNA is found in the nucleoplasm, and different from that seen in chick where heterochromatin is dispersed within the nucleoplasm nucleolus. This chimeric system provided reliable information on the fate and ontogeny of the engrafted cells.

Few years later, Rudolf Jaenisch developed a mouse chimeric system that allows the assessment of the developmental potential and migration of mouse neural crest in vivo. Primary neural crest cells isolated from C57BL/6 mouse embryos and microinjected in utero into neurulating E8.5 albino BALB/c embryos were shown to contribute efficiently to pigmentation in the host animal. The resulting neural crest chimeras showed, however, different coat pigmentation contribution-patterns depending on the genotype of the host embryos. Whereas BALB/c neural crest chimeras showed limited donor cell pigment contribution, restricted largely to the head and limbs, Kit^W-sh/Kit^W-sh mutant mice used as hosts for chimeras, displayed extensive pigmentation throughout, often exceeding 50% of the coat. The Kit^W-sh/Kit^W-sh murine model carries an inversion that spans a 2.8 Mbp segment proximal to the c-Kit locus that disrupts its regulatory sequences and leads to a deficit of melanocytes, with no change in viability or fertility. This mouse line allows for an empty melanoblast niche in which the transplanted NCCs can incorporate without competition from the host endogenous cell populations. In contrast to BALB/c chimeras, where the donor melanoblasts appeared to have migrated primarily in the characteristic dorsoventral direction, in Kit^W-sh/Kit^W-sh mutants the injected cells appeared to migrate into the longitudinal direction as well, as if the cells were spreading through an empty niche. This is consistent with the absence of a functional endogenous melanoblast population in Kit^W-sh/Kit^W-sh mutants, in contrast to BALB/c mice, which contain a full complement of melanocytes.

The human induced pluripotent stem cell (hiPSCs) technology provides patient specific pluripotent cells that carry all genetic alterations that contributed to the disorder and thus represent a genetically defined cell system to study the respective disease. The greatest promise of hiPSCs is its potential to study human diseases in the Petri dish. In this approach patient-derived cells are differentiated into the cell types, which is affected in the patient with the goal to uncover a disease relevant phenotype in the dish. This method was also applied for modeling neurocristopathies such as Familial Dysautonomia, by differentiating patient-derived cells hiPSCs to neural crest and its derivatives, which presented disease manifestation in vitro. However, numerous human diseases originate already in embryogenesis, i.e. are caused by disturbances of developmental processes, therefore such an approach cannot recapitulate the developmental aspects of a disease because the test cells are not incorporated into the developing embryo and do not participate in normal developmental processes. Thus, a major challenge of the “disease in the dish” approach is establishing model systems that, using patient-derived hiPSCs, allow for the investigation of human disease under long-term in vivo conditions.

Inspired by the ability to generate neural crest chimeras, we aspired to use the human pluripotent stem cells as a cellular source of human NCCs and to generate mouse-human neural crest chimeras. This platform would expand the potential of using human pluripotent stem cells for studying human neural crest development and disease in vivo. In our recent paper we have shown that human NCCS derived from human pluripotent stem cells when microinjected into post-implantation Kit^W-sh/Kit^W-sh mouse embryos could participate in normal embryonic development and provide functional neural crest contribution to the host murine model. We tracked the implanted NCCs, which had been GFP labeled, thru their migration paths, and found that the human cells exhibited similar migration patterns as would normally be found in mice. As a result, about 30% of the implanted embryos showed human NCCs during development, and later in adult mice pigmentation, which is similar to what we found in mouse-mouse neural crest chimeras.

Our results are one of the first evidences of contribution of human embryonic cell population with functional evidence in adult postnatal mammal. Interestingly, we have observed that the largest the evolutionary distance between the NCCs and the host mice, the less the contribution is noticed. While a widespread pigmentation was observed using primary mouse NCCs as donor cells, only localized pigmented hairs were found in Kit^W-sh/Kit^W-sh host mice, using rat NCCs and minimal for in vitro derived human NCCs. The limited level of contribution we observed using human NCCs, presumably represents the over 90 million years evolutionary distance between mouse and human, suggesting that the host environment might limit the maturation and differentiation of the injected human NCCs.

Our work serves as proof of concept and an important first step toward the goal of generating chimeric mice that carry disease-relevant human cells in the relevant tissue. Resulting mouse-human chimeras would fill an important gap in disease research, as existing models do not accurately mimic certain diseases and disease states. Cancer is frequently studied using xenografts, however this approach fails to provide insight into tumor initiation and progression. Moreover, complex diseases with long latencies, such as Alzheimer’s and Parkinson’s disease, can only be partially modeled using induced pluripotent stem cells in vitro. Mouse-human chimeras would be used to overcome these limitations and could be used for regenerative medicine as well.

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By Clare Buckley and Rachel Moore

One of the things that we find most challenging about working with whole vertebrate organisms is how we can tie ourselves in knots trying to unpick the function of a single component within such an intricate and interconnected web of proteins and signalling cascades. All too often knocking out (or down) a single gene produces a widespread phenotype, making it near impossible to differentiate between direct and indirect consequences. Either that, or redundancy within the system means that there is no phenotype at all. The recent morphant vs. mutant vs. CRISPR debate highlights this difficulty (Kok et al., 2015; Rossi et al., 2015; Schulte-Merker and Stainier, 2014).

The ‘chicken and egg’ conundrums of cell polarity

We were getting particularly frustrated with this problem while trying to work out how to test a feedback loop that we proposed following our finding of a novel mechanism of cell polarisation during zebrafish hindbrain lumen formation. Using live confocal imaging, we found that neural progenitor cells locate their centrosomes and assemble apical polarity complexes to whichever point they intersect the middle of the developing tissue, even if this is part-way along a cell length (Buckley et al., 2013).

This initiation of apical polarisation part way along the length of a cell, rather than at a cell extremity, was unexpected and raised the question of how these cells ‘know’ where to initiate apical polarity establishment. We proposed a polarisation feedback loop by which initial Pard3 puncta specify centrosomal location, causing the centrosome to organise the microtubule cytoskeleton around the tissue midline, which is in turn necessary to reinforce Pard3 delivery to this point.

Whilst watching the localisation of individual proteins during morphogenesis often tells us a lot about their function, directly testing this feedback loop will require us to manipulate one or more of its components. Rather than abrogating the function of e.g. Pard3, a much more powerful approach in this case would be to physically MOVE one of the components of the proposed feedback loop, to see which other components followed it. In order to do this, we needed a technique that would allow us to target specific proteins and to move them subcellularly with high temporal and spatial resolution.

The phytochrome system

We were mulling over this problem when we attended a conference on polarity at the Royal Society, where Orion Weiner presented his lab’s fantastic work using the Phytochrome system to manipulate subcellular protein localisation in cultured cells and yeast (Levskaya et al., 2009; Toettcher et al., 2013; Yang et al., 2013). For example, Levskaya et al. (2009) neatly illustrated that they could control lamellipodia extension in mammalian cells by targeting Rho-GTPases to specific sites in the cell membrane. Originally found in Arabidopsis, the phytochrome system consists of three components: the phytochrome B protein (PHYB); a basic-helix-loop-helix transcription factor called phytochrome interaction factor (PIF; PIF3 or PIF6); and an external chromophore, phycocyanobilin (PCB). PHYB and PIF are induced to bind in the presence of far-red light and to disassociate under infrared light. This interaction is dependent on PHYB being bound to the chromophore PCB (Ni et al., 1999).

The phytochrome system therefore had all of the characteristics we were looking for: spatiotemporal control that was not only fine-scale, but also reversible. It also had the advantage of utilising wavelengths (far-red and infrared) that we don’t often use for imaging cells, so we would be able to label and image our cells using standard green fluorophores. If only we could use this system in whole vertebrates!

Luckily for us, Orion and his lab share our view that collaboration is one of the keys to good science and Clare was able to visit his lab at UCSF on an EMBO short term fellowship to develop the phytochrome system so that it could be used in live zebrafish embryos. After a few months of cloning, swearing (relatively quietly) at the confocal microscope, fantastic interactions with the lab and plenty of Anchor beer, it finally worked! (Science being science, this occurred at 2am after some slightly louder swearing and with the aid of another late-working postdoc who helped Clare’s sleep-deprived brain switch the microscope on properly.)

The following year, back in Jon Clarke’s lab at King’s College London, we were able to develop the Phytochrome system further and use it in combination with live imaging of the developing zebrafish hindbrain, which resulted in our paper describing this method (Buckley et al., 2016).

Rapid and reversible

During the resubmission of our paper, the Phytochrome system was used to demonstrate nuclear protein import in superficially located zebrafish cells using whole-embryo illumination (Beyer et al., 2015). However, it had not yet been used in multicellular organisms for fine-scale spatiotemporal or reversible control of protein localization, or for accessing cells deeper within the tissue. Following optimization of the PHYB protein and of PCB chromophore purification and delivery, the phytochrome system worked well in zebrafish embryos. We injected a combination of a membrane-bound PHYB-mCherry-CAAX and cytoplasmic PIF6-EGFP mRNA along with PCB and measured how quickly the binding was turned on and off by analysing the intensity of EGFP fluorescence in the cytoplasm compared to the membrane. We found that the PHYB-PIF interaction provided us with rapid temporal control. Using the 633 nm laser on our confocal (far-red light induces PHYB-PIF interaction), PIF6-EGFP was recruited from the cytoplasm to the membrane with a time constant of 6.5 seconds. Conversely, PIF6-EGFP was released from the membrane into the cytoplasm at a time constant of 46.9 seconds under infrared light (which causes PHYB and PIF to dissociate) from a standard light LCD light source fitted with an infrared filter. This cycle of turning the binding off and on could be repeated several times without bleaching. And, crucially, we were successfully able to recruit the PIF6-EGFP construct to specific regions of the plasma membrane by restricting the far-red light (turning binding on) to a small region-of-interest whilst simultaneously bathing the entire sample in infrared light (turning binding off).

Beginning to address our original question – can we manipulate polarity protein location?

Of course, our original aim was to manipulate the polarity proteins from our putative feedback loop. To do this, we made a Pard3-PIF6-EGFP construct and confirmed that it showed a similar localisation to endogenous Pard3. We were able to direct its subcellular localisation into small regions-of-interest in the thin, flat epithelial cells of the enveloping layer, as well as in neuroepithelial cells in the developing neural tube.

Meanwhile, another project of Clare’s resulted in the birth of her daughter in April! Rachel heroically took up the mantle and produced some fantastic data showing that the specific enrichment of Pard3-PIF6-EGFP also resulted in the recruitment of the binding partner protein Pard6-MCherry to the same area, illustrating that Pard3 maintains its activity despite our manipulations. This demonstration of the spatiotemporal control of Pard3 activity is very exciting for us and opens up many possible experimental avenues. For example, during neural tube development, the inheritance of Pard3 by one daughter cell following mitosis is associated with neuronal fate (Alexandre et al., 2010; Dong et al., 2012). As a proof-of-principle experiment, we used the phytochrome system to force the Pard3-PIF6-EGFP into one daughter cell during a mitotic division. We think that this is an excellent example of the powerful experiments that will be possible using the phytochrome system.

Future work

There are a couple of further modifications that we think will make the phytochrome system even more robust. One obvious step forward would be to develop fish lines that express various components of the phytochrome system – PCB, for example, or the membrane-bound PHYB-mCherry-CAAX, are two that would be particularly useful.

Another important fact to remember is that we are not labelling – and therefore not disturbing – the endogenous Pard3 protein. To perfect this system we will need to either remove the endogenous protein before expressing the modified, PIF6-bound protein, or else insert the PIF6 sequence into the appropriate location close to Pard3. New techniques such as CRISPR should make the latter relatively straightforward.

We’re really looking forward to applying the phytochrome system to other questions in our lab as well. We also hope that it will be a useful tool for the developmental biology community. We envisage that such a subtle manipulation of proteins could be used to address a variety of problems, and we hope that it will be able to be applied to other model organisms. As such, we would encourage anyone who would like to try it out to contact us for constructs or a chat, and we look forward to reading about more experiments utilising the phytochrome system for developmental biology and beyond.

Alexandre, P., Reugels, A.M., Barker, D., Blanc, E., and Clarke, J.D. (2010). Neurons derive from the more apical daughter in asymmetric divisions in the zebrafish neural tube. Nature neuroscience 13, 673-679.

Beyer, H.M., Juillot, S., Herbst, K., Samodelov, S.L., Muller, K., Schamel, W.W., Romer, W., Schafer, E., Nagy, F., Strahle, U., et al. (2015). Red Light-Regulated Reversible Nuclear Localization of Proteins in Mammalian Cells and Zebrafish. ACS synthetic biology 4, 951-958.

Buckley, Clare E., Moore, Rachel E., Reade, A., Goldberg, Anna R., Weiner, Orion D., and Clarke, Jonathan D.W. (2016). Reversible Optogenetic Control of Subcellular Protein Localization in a Live Vertebrate Embryo. Developmental Cell 36, 117-126.

Buckley, C.E., Ren, X., Ward, L.C., Girdler, G.C., Araya, C., Green, M.J., Clark, B.S., Link, B.A., and Clarke, J.D. (2013). Mirror-symmetric microtubule assembly and cell interactions drive lumen formation in the zebrafish neural rod. The EMBO journal 32, 30-44.

Dong, Z., Yang, N., Yeo, S.Y., Chitnis, A., and Guo, S. (2012). Intralineage directional Notch signaling regulates self-renewal and differentiation of asymmetrically dividing radial glia. Neuron 74, 65-78.

Kok, Fatma O., Shin, M., Ni, C.-W., Gupta, A., Grosse, Ann S., van Impel, A., Kirchmaier, Bettina C., Peterson-Maduro, J., Kourkoulis, G., Male, I., et al. (2015). Reverse Genetic Screening Reveals Poor Correlation between Morpholino-Induced and Mutant Phenotypes in Zebrafish. Developmental Cell 32, 97-108.

Levskaya, A., Weiner, O.D., Lim, W.A., and Voigt, C.A. (2009). Spatiotemporal control of cell signalling using a light-switchable protein interaction. Nature 461, 997-1001.

Ni, M., Tepperman, J.M., and Quail, P.H. (1999). Binding of phytochrome B to its nuclear signalling partner PIF3 is reversibly induced by light. Nature 400, 781-784.

Rossi, A., Kontarakis, Z., Gerri, C., Nolte, H., Holper, S., Kruger, M., and Stainier, D.Y.R. (2015). Genetic compensation induced by deleterious mutations but not gene knockdowns. Nature 524, 230-233.

Schulte-Merker, S., and Stainier, D.Y.R. (2014). Out with the old, in with the new: reassessing morpholino knockdowns in light of genome editing technology. Development 141, 3103-3104.

Toettcher, J.E., Weiner, O.D., and Lim, W.A. (2013). Using optogenetics to interrogate the dynamic control of signal transmission by the Ras/Erk module. Cell 155, 1422-1434.

Yang, X., Jost, A.P., Weiner, O.D., and Tang, C. (2013). A light-inducible organelle-targeting system for dynamically activating and inactivating signaling in budding yeast. Molecular biology of the cell 24, 2419-2430.

(4 votes)

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A postdoctoral position is available to study the cellular basis of morphogenesis in vertebrate craniofacial development. This work will integrate mouse molecular genetic approaches with live cell imaging, cell biology and biochemistry to study signaling mechanisms in development, and how this signaling goes wrong in congenital disease (e.g. PLos Biology 2015 13(4): e1002122). The position is in the laboratory of Jeff Bush (bush.ucsf.edu), in the UCSF Department of Cell and Tissue Biology and Program in Craniofacial Biology. The laboratory is located at the UCSF Parnassus Heights campus, in the center of San Francisco. UCSF offers an outstanding developmental biology community and a supportive working environment.

Candidates with a Ph.D. degree in a biological science and research experience in live cell or live embryo imaging, molecular biology, genetics, or biochemistry should submit a C.V. and names of at least 2 references via email to: jeffrey.bush@ucsf.edu

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Morphogenesis, the shaping of tissues and organs, is driven by a series of events that proceed in a coordinated manner, both spatially and temporally. Such events include changes in cell shape, cell adhesion, and cell migration, which happen at a precise developmental stage in single cells or cell collectives. The detailed study of morphogenetic processes relies upon the ability to perturb these localized changes in the otherwise intact organism, ideally with sub-cellular precision and on a time-scale of seconds. However, this is impossible to achieve with traditional genetic or chemical tools.

When, in 2011, I started my PhD in the De Renzis group at EMBL, optogenetics had just been elected “Method of the Year” by Nature Methods¹ and was included in Science’s “Breakthroughs of the Decade”². Since its early approaches at the beginning of 2000s^3-5, optogenetics has greatly facilitated the study of neuronal circuits in the brain. In these pioneering experiments, mice were made to express light-sensitive ion channels in selected populations of neurons, thus allowing control over brain activity with a pulse of laser light. By the end of 2000s, a few studies reported using optogenetics also in non-excitable cells^6-8. Despite these advances, being able to control morphogenetic movements with light still represented a challenge.

My longstanding interest in biotechnology led me to propose co-opting optogenetics to modulate cell behaviour during Drosophila morphogenesis. Data from our lab had shown that a particular plasma membrane lipid, phosphatidylinositol-4,5-bisphosphate (PI(4,5)P₂), is essential for cell morphogenesis in the Drosophila embryo⁹. Indeed, PI(4,5)P₂ is known to act as a molecular scaffold for actin-binding proteins, thereby regulating actin polymerization at the cell cortex. PI(4,5)P₂ depletion from the plasma membrane is mediated by a class of enzymes known as inositol-polyphosphate-5-phosphatases (5-phosphatases). I reasoned that by modulating PI(4,5)P₂ metabolism at the plasma membrane, I should be able to deplete actin from the cell cortex. Because cortical actin is key for many biological processes ranging from cell contractility to cell division, this tool would allow the control of a wide spectrum of morphogenetic events. The not-so-trivial question was how to do that using light.

Just a couple of years before I started my PhD, a few research groups had developed optogenetic approaches to specifically control protein-protein interaction in cell culture^10-12. After considering the functional features of these tools, I decided to use the Cry2-CIB1 system¹², as it can be rapidly activated with blue light and does not need the addition of an exogenous chromophore. The Cry2-CIB1 system is composed of two photosensitive modules: Cryptochrome 2 (Cry2) and CIB1, which interact only in the presence of blue light (458-488 nm). This system had already proved suitable to control phosphoinositide metabolism in cultured cells¹³. In the optogenetic approach I set up, a GFP-tagged CIB1 localizes to the cells’ plasma membrane, whereas the 5-phosphatase, which is fused to Cry2 and mCherry, stays in the cytosol. The fluorescent tags would allow me to monitor the protein location in vivo. To test whether I could use this system to modulate morphogenetic movements, I decided to focus on ventral furrow formation, the invagination of the Drosophila mesodermal tissue, as this process is highly dependent on actin dynamics and cell contractility. During ventral furrow formation, a stripe of ~1000 ventral cells constrict their apical surfaces and fold inwards as a tube. The core idea is that in the dark, the 5-phosphatase would remain cytosolic, and morphogenesis would proceed normally (Figure 1A). In the presence of blue light, the 5-phosphatase would relocate to the plasma membrane, where it would deplete PI(4,5)P₂ and, in turn, actin from the cell cortex. This should result in the inhibition of cell contractility and a blockade of ventral furrow formation (Figure 1B).

When I started to test this optogenetic approach, I first looked at the effects of blue light illumination on the 5-phosphatase localization, and on PI(4,5)P₂ and actin levels. Upon a 1 s pulse of 488 nm light, the 5-phosphatase relocated to the plasma membrane. The relocalization of the 5-phosphatase resulted in a sharp reduction in PI(4,5)P₂ and cortical actin levels. I therefore went on to characterize the effects of PI(4,5)P₂ and actin depletion on cell contractility during ventral furrow formation. Illumination of the entire embryo with a 488 nm (single photon) laser resulted in the inhibition of both apical constriction and tissue folding. After this encouraging result, I sought to achieve spatial specificity in addition to temporal precision by photo-activating smaller groups of cells. However, light scattering within the tissue resulted in unwanted photo-activation of cells neighboring the illuminated areas, making the single photon approach impracticable. By discussing with in-house microscopy experts, I found out that this limitation could be overcome by using two-photon microscopy. Indeed, the requirement for near-simultaneous absorption of two photons makes it very likely that photo-activation remains limited to the area where the laser is focused. I therefore set up a protocol to achieve optimal levels of photo-activation with a two-photon laser (950 nm). To my delight, photo-activation could be achieved at the seconds time-scale and allowed high spatial precision, namely single cell resolution.

Having established this powerful approach, I tackled two interesting questions regarding ventral furrow formation. In particular, I tested whether apical constriction in ventral cells is required only to initiate the process of tissue bending, or whether it is necessary throughout the process to achieve complete folding of the tissue. By modulating apical constriction only in ventral cells, I could show that apical constriction is necessary not only to kick off, but also to sustain tissue folding.

Another open question concerned the way individual cells constrict during ventral furrow formation. When ventral cells reduce their apical surface, they do so in an asymmetric fashion: cells shrink along the embryo’s medial-lateral axis and remain elongated along the anterior-posterior (a-p) axis. This asymmetry in cell constriction, known as a-p anisotropy, is thought to be the result of higher tension along the a-p axis than along the medial-lateral axis. However, the origin of this tension was unknown. To address this question, I altered the geometry of the primordium by inhibiting cell contractility in two areas at the anterior and posterior end of the ventral furrow tissue. Then, I checked whether this resulted in any change in a-p anisotropy in cells that were left able to constrict (i.e. not illuminated with blue light). I could show that the degree of a-p anisotropy in non-illuminated cells was higher if the two illuminated areas were further apart than if they were close together along the a-p axis (Figure 2). In other words, if the rectangular geometry of the constricting tissue was preserved cells constricted in an asymmetric way. Instead, if the constricting patch of cells was squared-shaped, constriction was more symmetric, and resulted in roundish cells. This suggests that the geometry of the ventral furrow tissue impacts on the way individual cells constrict.

As I collected increasing amount of data, I needed to transform a lot of visual information into meaningful numbers, so I teamed up with Joseph Barry, who at that time was a postdoc in Wolfgang Huber’s group at EMBL. Joe did a great job of developing algorithms to quantify cell features, such as area and a-p anisotropy. For this project, I heavily relied on the EMBL Advanced Light Microscopy Facility, the state-of-the-art equipment available in the Developmental Biology Unit, and of course the valuable input of my supervisor Stefano De Renzis. Shortly after publication in Developmental Cell¹⁴, our optogenetic approach generated the interest of many research groups around the world, as apical constriction is a highly conserved cellular mechanism during morphogenesis. However, besides apical constriction, cell contractility drives a multitude of cell behaviors, from cytokinesis to cell migration. Therefore, I expect that our approach will be useful to the community for addressing intriguing questions about animal development.

In the meanwhile, optogenetics has enabled the modulation of many other cellular activities, including organelle transport, cell-cell signalling, apoptosis, and gene expression^15-20. The translation of these tools to living organisms will undoubtedly provide new insights into the mechanisms by which embryos grow and develop. As a developmental biologist and recent PhD graduate, I am very much looking forward to seeing where lights will guide us.

^{1. Method of the Year 2010. Nat Meth 8, 1-1, doi:10.1038/nmeth.f.321 (2011). ↩}

^{2. Insights of the decade. Stepping away from the trees for a look at the forest. Introduction. Science (New York, N.Y.) 330, 1612-1613, doi:10.1126/science.330.6011.1612 (2010). ↩}

^{3. Zemelman, B. V., Lee, G. A., Ng, M. & Miesenbock, G. Selective photostimulation of genetically chARGed neurons. Neuron 33, 15-22 (2002). ↩}

^{4. Banghart, M., Borges, K., Isacoff, E., Trauner, D. & Kramer, R. H. Light-activated ion channels for remote control of neuronal firing. Nat Neurosci 7, 1381-1386, doi:10.1038/nn1356 (2004). ↩}

^{5. Boyden, E. S., Zhang, F., Bamberg, E., Nagel, G. & Deisseroth, K. Millisecond-timescale, genetically targeted optical control of neural activity. Nat Neurosci 8, 1263-1268 (2005). ↩}

^{6. Wu, Y. I. et al. A genetically encoded photoactivatable Rac controls the motility of living cells. Nature 461, 104-108, doi:10.1038/nature08241 (2009). ↩}

^{7. Wang, X., He, L., Wu, Y. I., Hahn, K. M. & Montell, D. J. Light-mediated activation reveals a key role for Rac in collective guidance of cell movement in vivo. Nature cell biology 12, 591-597, doi:10.1038/ncb2061 (2010). ↩}

^{8. Toettcher, J. E., Gong, D., Lim, W. A. & Weiner, O. D. Light-based feedback for controlling intracellular signaling dynamics. Nature methods 8, 837-839, doi:10.1038/nmeth.1700 (2011). ↩}

^{9. Reversi, A., Loeser, E., Subramanian, D., Schultz, C. & De Renzis, S. Plasma membrane phosphoinositide balance regulates cell shape during Drosophila embryo morphogenesis. The Journal of cell biology 205, 395-408, doi:10.1083/jcb.201309079 (2014). ↩}

^{10. Levskaya, A., Weiner, O. D., Lim, W. A. & Voigt, C. A. Spatiotemporal control of cell signalling using a light-switchable protein interaction. Nature 461, 997-1001, doi:10.1038/nature08446 (2009).↩}

^{11. Yazawa, M., Sadaghiani, A. M., Hsueh, B. & Dolmetsch, R. E. Induction of protein-protein interactions in live cells using light. Nature biotechnology 27, 941-945, doi:10.1038/nbt.1569 (2009).↩}

^{12. Kennedy, M. J. et al. Rapid blue-light-mediated induction of protein interactions in living cells. Nature methods 7, 973-975, doi:10.1038/nmeth.1524 (2010).↩}

^{13. Idevall-Hagren, O., Dickson, E. J., Hille, B., Toomre, D. K. & De Camilli, P. Optogenetic control of phosphoinositide metabolism. Proceedings of the National Academy of Sciences of the United States of America 109, E2316-2323, doi:10.1073/pnas.1211305109 (2012).↩}

^{14. Guglielmi, G., Barry, J. D., Huber, W. & De Renzis, S. An Optogenetic Method to Modulate Cell Contractility during Tissue Morphogenesis. Developmental cell, doi:10.1016/j.devcel.2015.10.020 (2015).↩}

^{15. Mills, E., Chen, X., Pham, E., Wong, S. & Truong, K. Engineering a photoactivated caspase-7 for rapid induction of apoptosis. ACS synthetic biology 1, 75-82, doi:10.1021/sb200008j (2012). ↩}

^{16. Liu, H., Gomez, G., Lin, S., Lin, S. & Lin, C. Optogenetic control of transcription in zebrafish. PloS one 7, e50738, doi:10.1371/journal.pone.0050738 (2012).↩}

^{17. Toettcher, J. E., Weiner, O. D. & Lim, W. A. Using optogenetics to interrogate the dynamic control of signal transmission by the Ras/Erk module. Cell 155, 1422-1434, doi:10.1016/j.cell.2013.11.004 (2013).↩}

^{18. Grusch, M. et al. Spatio-temporally precise activation of engineered receptor tyrosine kinases by light. The EMBO journal 33, 1713-1726, doi:10.15252/embj.201387695 (2014).↩}

^{19. Motta-Mena, L. B. et al. An optogenetic gene expression system with rapid activation and deactivation kinetics. Nature chemical biology 10, 196-202, doi:10.1038/nchembio.1430 (2014).↩}

^{20. van Bergeijk, P., Adrian, M., Hoogenraad, C. C. & Kapitein, L. C. Optogenetic control of organelle transport and positioning. Nature 518, 111-114, doi:10.1038/nature14128 (2015).↩}

(5 votes)

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Last week saw ASAPbio, a meeting that discussed the role that preprints can play in the life sciences (for a an introduction to preprints check out this video or this wikipedia page).  Those of you on twitter will have followed the #ASAPbio discussion with interest,  and the footage of the conference is now available online. What is your experience: have you deposited your manuscript on a preprint service like bioRxiv? If not, have you considered doing so, and what would persuade/deter you? This month we are asking:

What is the value of preprint servers in Biology?

Share your thoughts by leaving a comment below! You can comment anonymously if you prefer. We are also collating answers on social media via this Storify. And if you have any ideas for future questions please drop us an email!

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In 2014, Development organised a very successful meeting on how the use of stem cell technologies can inform our understanding of human development (you can read about it here or watch the movie below). The next edition of this meeting will take place in the USA this September and applications are now open! The deadline for applications is the 15th of June, but we encourage you to apply early to avoid disappointment. Just click the image above or follow this link.

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ISBN 9789814678537 – 28USD/18GBP

The idea of human germline genetic modification is too close for comfort right now. However, society in general does not seem to realise the proximity of this threat or the technical basis of this threat, making the publishing of Paul’s book ‘GMO Sapiens – The Life-Changing Science of Designer Babies’ timely. This book is 250 pages of a digestible history of genetic modification and technologies and opinions that are leading to the very real and urgent threat of heritable human genetic modification.

In reading this book it should be noted from the start that while the validation of many that modification of the human germline would be useful for eliminating human disease, in the light of existing, safer and more efficient PGD (pre-implantation genetic diagnosis ) technologies (Chapter 5), this argument seems to front a human desire for modifications towards predicted ‘improvement’ of humans, Human+ if you want to go for the transhumanist view (Chapter 7). Personally, I do not believe there is a valid argument for genetic modification of the nuclear genome in the human germline (in light of PGD). To hide ‘positive eugenics’ (Chapter 7) behind a veil of ‘eliminating human disease’ is frankly wrong. Expert interviews in this book from renowned scientists shamelessly expressing their desire to ‘improve’ humans is scary. However, it was mentioned early that creating panic or scaring people is not the goal of this book, but rather to inform and energize people to become part of the discussion about this new inevitable reality. Reading this book is going to catch you up with the best of the best.

If we stick to arguments and perspectives on the heritable genetic ‘improvement’ of humans, Paul provides a thorough and balanced view. Who should choose what is better? Do we know enough about the human genome to safely predict ‘better’? Will better be for the individual or for society as a whole? All of these views and more are discussed in the book giving time to both sides. There are however a lot of question marks, which reasonable people will approach with caution. But it is not the reasonable who will be pioneering clinical germline modification. It is a fact that someone will heritably modify the human genome (someone has – Chapter 4 and Chapter 9) and use this to enhance perceived positive traits. How will we react to this? How will we manage the unseen consequences.

However, once the mistakes have been made, the technology refined and the benefits become clear, would I say no? Paul presents a story where you have yourself wondering what choices you would make if presented with a list of options; Should I choose that my children are resistant to diseases? Should I choose that my children have reasonably sizes physical features so they do not get teased at school? In a society where these options were available as a reproductive shopping list, what parent would not choose what is ‘best’ for their child? Are we aiming towards human agriculture?

There may be someone in the world right now miscarrying or having an abortion from a defective genetically modified human embryo. This urgency and a thread of caution is present throughout the book. But what I find troubling, exciting but scary, is that I find myself agreeing with an undertone, I do not support human germline genetic modification but with all the new information and perspectives available to me I have found myself questioning my own views and will be watching any developments with a fascinated interest I would rather not admit to.

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https://www.jobs.manchester.ac.uk/displayjob.aspx?jobid=10973

We are seeking an enthusiastic and outstanding postdoctoral researcher to join a multidisciplinary team led by Prof. Chris Thompson.  This project aims to address the role and regulation of heterogeneity in development using Dictyostelium discoideum as a model system. Dictyosteliumpermits a uniquely powerful combination of approaches to be applied (e.g. imaging, informatics, genetics) and thus provides an opportunity and generate the first integrative ‘top to bottom’ understanding of how heterogeneity, stochastic differentiation and cell sorting result in robust developmental patterning.

You will address the fundamental biological questions regarding the role and regulation of heterogeneity in development you will use Dictyostelium discoideum as a model system. Dictyosteliumpermits a uniquely powerful combination of approaches to be applied.  Firstly, developmental patterning in Dictyostelium is based on ‘salt and pepper’ differentiation followed by sorting out, and therefore heterogeneity has been proposed to play a pivotal role.  Secondly, Dictyostelium is amenable to forward and reverse genetic manipulation, is easily and rapidly grown in the lab to biochemical scales, whilst its relatively small number of defined cell types can be tracked in vivo by live cell imaging during development. Dictyostelium therefore provides an opportunity and generate the first integrative ‘top to bottom’ understanding of how heterogeneity, stochastic differentiation and cell sorting result in robust developmental patterning.

You will use your extensive experience in bioinformatics, computational biology, molecular biology, genetics, cell biology or live cell imaging techniques to determine the molecular basis and gene networks that regulate heterogeneity. These different approaches are highly complementary and your ability to integrate these approaches is crucial. Consequently, multidisciplinary training (especially in computational and wet lab skills) is essential. You should currently hold or be about to obtain a PhD in a relevant field.

Although you will be based in Manchester, several short visits to collaborators will be required for data analysis and project development.

The post funded by the Wellcome Trust and is available for up to 3 years.

Successful candidates will be subject to pre-employment screening carried out on our behalf by a third party. The offer of employment will be dependent on the successful candidate passing that screening. Whilst you will be required to provide express consent at a later stage, by continuing with your application now you acknowledge that you are aware that such screening will take place, and agree to take part in the process.

The School of Life Sciences is committed to promoting equality and diversity, including the Athena SWAN charter for promoting women’s careers in STEMM subjects (science, technology, engineering, mathematics and medicine) in higher education. The School received a Silver Award in 2009 for their commitment to the representation of women in the workplace and we particularly welcome applications from women for this post. Appointment will always be made on merit. For further information, please visit: http://www.wils.ls.manchester.ac.uk/athenaswanawards/

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Researchers at IRB Barcelona and CSIC discover a mechanism through which the cells of an organism interact with their extracellular matrix

The cells of an organism interact not only with each other but with the extracellular matrix that surrounds them. Increasing evidence is unveiling the relevance of this structure—which is secreted by the cells themselves— for the correct function of the organism and also for the development of various diseases.

A new study published in eLife and headed by Jordi Casanova and Sofía J. Araújo, both scientists at the Institute for Research in Biomedicine (IRB Barcelona) and the Instituto de Biología Molecular de Barcelona (IBMB-CSIC), describes a cell communication mechanism that allows the organisation of the extracellular matrix and how this structure affects cells through a feedback system.

For this study, the team of researchers used the fruit fly Drosophila melanogaster—a particular useful model for biomedical research. The study focused on the tracheal system, tubes that are analogous to the function of the human respiratory apparatus. This system has an extracellular matrix that covers the inside of the trachea, forming a structure that is comparable to the hose of a vacuum cleaner. Until now, it was believed that this matrix served only a structural purpose, preventing the tube from collapsing, but the team of scientists has demonstrated that it also regulates the cells that form it.

In 1929, the Canadian biologist W. R. Thompson published a study describing the tracheal system and its structure. Although he was able to describe it, he was unable to explain how it formed. This new study now provides an explanation of this 80-year enigma.

“The biological context of these cells modifies not only their behaviour but also their internal structure,” comments Casanova. “When we modify only the extracellular matrix, the cytoskeleton is also altered.”

“It is a two-fold mechanism,” says Sofía Araújo. “First actin filaments, a very important component of the cytoskeleton, serve as a mould for the deposition of the chitin of the matrix. Next, the matrix itself stabilises the cytoskeleton, anchoring actin in place.” The scientists propose that Src42A—a protein that belongs to the family of kinases that regulates the structure of the actin filament—is one of the main contributors to this system.

Casanova considers that the study explains one of the many mechanisms that allow communication between the extracellular matrix and cells. “The way in which cells communicate has been conserved over evolution: we are sure that this process will be discovered in other organisms. In our lab, we address how such communication allows cells to arrange themselves in such a way as to form tissues.”

The interaction between the cell and its extracellular matrix is also very important in inflammatory and cancer processes. “Tumour cells often take advantage of existing mechanisms, such as the one we have described, to cause havoc. The unravelling of these mechanisms may provide us with new tools to study diseases,” concludes Casanova.

Reference article:

A feedback mechanism converts individual cell features into a supracellular ECM structure in Drosophila trachea

Arzu Öztürk-Çolak, Bernard Moussian, Sofia J. Araújo and Jordi Casanova

eLife (2016): doi: 10.7554/eLife.09373.001

This article was first published on the 22nd of February 2016 in the news section of the IRB Barcelona website

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