Here are the highlights from the current issue of Development:

Mak(or)in’ the switch to adulthood

The juvenile-to-adult (J/A) transition of many animals, from worms to humans, is regulated by the highly conserved RNA-binding protein LIN-28. In this and other contexts – including stem cell renewal versus differentiation decisions – LIN-28 acts to suppress the production of the microRNA let-7, which in turn inhibits a suite of downstream genes, most notably the translational regulator LIN-41. Now, using the J/A transition of C. elegans as a model, David Fitch and colleagues (799) identify a new player in this axis, the Makorin orthologue LEP-2. lep-2 mutant adults display a number of juvenile characteristics, including failure of male tail tip retraction, continued moulting into adulthood and defective male sexual behaviour. The authors provide evidence that LEP-2 acts to promote degradation of LIN-28 in larval stages, which is necessary for the J/A transition. The underlying molecular mechanism has yet to be resolved: as a putative E3 ubiquitin ligase, LEP-2 might directly target LIN-28 for degradation or it may act indirectly. Given the conservation of the Makorin family, along with data implicating mammalian Makorins in cell state transitions and in the timing of puberty onset, it is possible that this Makorin/LIN-28 interaction could control developmental switches in multiple contexts.

Maintaining and reprogramming sexual identity

Gonadal sexual identity is determined during development, but must also be maintained in adulthood. In Drosophila, the transcription factor Chinmo has been identified as a key regulator of male identity in the adult testis: it promotes expression of the male sex determinant Dsx^M (the homologue of which, DMRT1, is a key regulator of male identity in mouse testes). In chinmo mutants, somatic stem cells of the adult testis adopt female fate. Now (754), Erika Matunis and co-workers show that Chinmo is not only necessary but also sufficient to promote male fate. Overexpression of chinmo in adult ovarian somatic cells leads to severe oogenesis phenotypes. Marker expression analysis suggests thatchinmo-overexpressing somatic cells lose female identity and gain male fate. Strikingly, this also appears to affect the sexual fate of the germ cells: a proportion of ovary germ cells start to express male markers upon somatic chinmo expression. Unlike in testis, Chinmo in the ovary does not appear to promote Dsx^M expression, implying that there must be alternative mechanisms for masculinisation of somatic cells. Although these mechanisms have yet to be uncovered, these data provide strong evidence that sexual identity of the Drosophila adult ovary can be reprogrammed, and that sexual fate must be actively maintained throughout life.

At the heart of histone methylation

Mutations in the histone methyltransferase KMT2D are associated with Kabuki Syndrome – a haploinsufficient congenital, multi-organ syndrome that frequently includes severe heart defects. However, the role of KMT2D in the heart has not been analysed in detail. Benoit Bruneau and colleagues now address this (810), generating several conditional Kmt2d mouse mutants and analysing them both phenotypically and using genome-wide approaches. They find that cardiac deletion of Kmt2d causes embryonic lethality with defects in heart morphology and cardiomyocyte proliferation. Global gene expression analysis demonstrates dysregulation of genes associated with cell cycle regulation, ion homeostasis and hypoxia signalling. Functionally, ventricular calcium handling appears impaired. KMT2D is involved in H3K4 mono- and di-methylation; consistent with this, ChIP-Seq data demonstrate that Kmt2d depletion causes loss of H3K4me1 and me2 at specific loci. By correlating these data with the RNA-seq profiles and ChIP-Exo data for KMT2D chromatin binding, the authors are able to identify a small number of high-confidence targets for KMT2D, functions of which are consistent with the phenotypes observed upon Kmt2d deletion. As well as shedding light on the important role of KMT2D in mouse heart development, these data may have implications for the aetiology of the heart defects observed in Kabuki syndrome.

TFAP2C: a key controller of placental growth

In mammals, proper placental development is essential for growth and viability of the embryo. The transcription factor TFAP2C is known to be important for specification and maintenance of trophoblast stem cells (placental progenitors), but whether this factor also plays roles at later stages of placental development is less well understood. On 787, Hubert Schorle and co-workers provide insights into the role of TFAP2C in a subset of placental progenitors, the TPBPA-expressing population that forms the junctional zone of the placenta. Loss of Tfap2c from this population leads to growth defects in the junctional zone, with reduced numbers of TPBPA⁺ cell-derived trophoblasts. Microarray analysis and follow-up experiments provide evidence that TFAP2C controls several key aspects of placental development: it inhibits Cdkn1a, a cell cycle inhibitor; promotes expression and activation of Akt to regulate glycogen synthesis; and promotes MAPK pathway activity – important for trophoblast proliferation and differentiation – by repressing the Dusp family of MAPK inhibitors. Importantly, this conditional mouse mutant provides a model for intrauterine growth retardation, as mutant embryos show lower foetal and birth weight. Preliminary data in a human trophoblast cell model suggests that this important role of TFAP2C may be conserved.

PLUS:

Building and re-building the heart by cardiomyocyte proliferation

Dissecting the cellular and molecular mechanisms that promote cardiomyocyte proliferation throughout life, deciphering why proliferative capacity normally dissipates in adult mammals and deriving means to boost this capacity, are primary goals in cardiovascular research. Here, Matthew Foglia and Kenneth Poss discuss the cellular and molecular mechanisms that control cardiomyocyte proliferation, during both heart development and regeneration across various species. See the Review article on p. 729

A comparative view of regenerative neurogenesis in vertebrates

Alessandro Alunni and Laure Bally-Cuif summarize the striking similarities in the essential molecular and cellular properties of adult neural stem cells between different vertebrate species, both under physiological and reparative conditions. They also discuss differences in the reparative process across evolution and how the study of non-mammalian models can provide insights into both basic neural stem cell properties and stimulatory cues shared between vertebrates. See the Review on p. 741

Featured movie

This movie shows the mating behaviour of wild type male C. elegans. This behaviour is disrupted in mutants of lep-2, a new heterochronic gene identified by Fitch and colleagues. Read the paper here.

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DMM is looking for an enthusiastic intern who wishes to gain experience in science publishing.

Joining an experienced and successful team, including Academic Editor-in-Chief Monica Justice, the internship offers an ideal opportunity to gain in-depth experience on a growing Open Access journal in the exciting and fast-moving field of translational research. DMM publishes primary research articles and a well-regarded front section of reviews and topical comment. The intern will work alongside an established publishing team in our Cambridge offices.

Because the journal serves both basic biomedical researchers and clinicians, applicants will have a PhD or MD, ideally with some relevant research experience, and a broad knowledge of model organisms and disease issues.

The intern’s core responsibilities will include:

• Representation of the journal at international scientific conferences, Company workshops and within the wider scientific community, with a view to promoting the journal and commissioning new front-section content.

• Conducting interviews with high-profile scientists in the biomedical arena.
• Support for the Academic Editors in their assessment and handling of primary research articles.

• Outlining a strategy for journal social media activities and writing press releases.

• Involvement in the journal’s development and marketing activities.
• Contribute to special editorial projects on the journal.

Essential requirements for the job are enthusiasm, commitment, judgement, integrity and a mature attitude. Candidates should have excellent interpersonal skills and confidence, excellent oral and written communication skills, and a broad interest in research and the research community. They should also be willing to travel. Previous editorial experience is not required.

The internship will last for 9 months at a salary of £15,000 pro rata.

The Company of Biologists is based in attractive modern offices on the outskirts of Cambridge, UK.

The Company of Biologists exists to support biologists and inspire advances in biology. At the heart of what we do are our five specialist journals – Disease Models & Mechanisms, Development, Journal of Cell Science, Journal of Experimental Biology, and Biology Open – two of them fully open access. All are edited by expert researchers in the field, and all articles are subjected to rigorous peer review. We take great pride in the experience of our editorial team and the quality of the work we publish. We believe that the profits from publishing the hard work of biologists should support scientific discovery and help develop future scientists. Our grants help support societies, meetings and individuals. Our workshops and meetings give the opportunity to network and collaborate.

Applicants should send a CV by email to hr@biologists.com along with a covering letter that summarises their relevant skills and experience, and why they are enthusiastic about this opportunity. Candidates must be able to demonstrate their entitlement to work in the UK.

Applications should be made as soon as possible and by March 21st 2016.

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Research

– Two different applications of optogenetics were highlighted on the Node this month. Giorgia wrote about how optogenetics can be used to control morphogenesis in Drosophila. Meanwhile, Clare and Rachel described how they adapted the phytochrome system to live zebrafish embryos, allowing proteins to be moved within a live embryo using light!

– What is the relationship between ESCs and cells in the mouse embryo? And how does naïve pluripotency differ in mouse and primate species? Thorsten posted about his latest paper and introduced a mouse and marmoset pathway expression atlas for early development.

– Carine and Nadine presented a new method to detect and quantify single RNAs at cellular resolution in zebrafish embryos.

– Malkiel presented mouse-human neural crest chimeras, and the applications of this system for studying human neural crest development and disease in vivo.

– And a 1898 paper by T. H. Morgan on planarian regeneration is the focus of our latest ‘Forgotten Classic’, following a recommendation by Alejandro Sánchez Alvarado.

A day in the life

This month saw two new additions to our ongoing series!

– Helena, who was doing her PIPS internship with us until recently, wrote about what it is like to do cell culture and work with ESCs.

– And the McGregor lab told us about what is like using house spiders to study developmental biology and development!

Discussion

– Have you ever deposited your paper in a pre print server like bioRxiv? What would persuade you to? Share your thoughts with the latest Question of the Month!

– Why are international collaborations important? James wrote about his visit to India to launch a new EMBO partnership.

– And we reposted a best practice guide to combating irreproducibility in mouse research, originally published in Disease Models & Mechanisms.

Also on the Node

– What’s the common thread between growth and regeneration? Katherine shared a report from a recent Keystone meeting on this topic.

– Are you a fly pusher? Mario developed a free software that will help you keep track of your genetic crosses!

– Two book reviews featured on the Node this month. Helena reviewed the book “Decoding the Language of Genetics” by David Botstein, while Heather considered the brave new world of human germline genetic modification by reading “GMO Sapiens” by Paul Knoepfler.

– and Karin visited the Huisken lab in Dresden to image zebrafish development using panoramic SPIM, sponsored by a Development Travelling Fellowship.

Would you like to run the Node? Last month we also announced that we are recruiting a new Node community manager! The deadline for applications is the 14th of March.

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Some time ago I wrote about a webpage I’ve created to manage genetic fly crosses.

In that past post, I’ve promised a new version of the software, and I’m happy to announce it here.

The software can be reached under the address http://drosophila.me. Many many things changed from the previous version, some features:

• User accounts
• Your crosses are saved by default, you can use the software as a sort of digital genetic notebook
• Crosses can be shared with colleagues (or with a qr code on a poster during a conference)
• Cross history can be followed over generations
• User can establish an own stock list and cross directly from it
• Several visualisations of a cross: punnett square, genotypes, curly bracket view
• Smartphone friendly

A web app as complex as drosophila.me is quite a code monster, and as I’m the only developer (for now), there might still be some glitches. I usually can solve them quickly, so let me know about it :)

I hope this tool will be useful for the community.

Welcome screen

Cross notebook

Genotypes view

Punnett square view

Curly bracket view

Stock list

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Places are still available for early career scientists interested in attending The Company of Biologists workshop ‘Metabolism in Development and Disease’. This is a unique opportunity to interact with the leaders in this field in a small and informal meeting. Apply until the 16th of March! For more information follow this link.

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Professors Graham Williams and Duncan Bassett have 6 full-time posts (5 years each) and 2 shared appointments (with Dr Jacques Behmoaras, Medical Research Council, 4 years each) to fill. You will be joining an established laboratory with state-of-the-art equipment and a strong track record in molecular endocrinology. The lab is a founder member of the Origins of Bone and Cartilage Disease (OBCD) international phenotyping consortium. Please see our website for this exciting opportunity to join our cutting-edge research program based at Imperial College London.

http://boneandcartilage.com/opportunities.html

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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).↩}

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