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Imaging specialist

Posted by , on 28 June 2018

Closing Date: 15 March 2021

 

A staff research position for an imaging specialist is available in the Parichy lab at University of Virginia. The laboratory focuses on cellular interactions and morphogenetic behaviors, with particular emphasis on post-embryonic neural crest derivatives including pigment cells.

The successful applicant will contribute to on-going studies, will have opportunities to design and pursue new projects, and will oversee microscopy and imaging infrastructure.

The laboratory is equipped with several instruments for high resolution imaging including:

  • Zeiss LSM 880 multi-photon laser scanning microscope with Fast Airyscan for super-resolution time-lapse
  • Zeiss LSM 800 laser scanning microscope with Airyscan for super-resolution
  • Zeiss AxioObserver with Yokogawa spinning disk
  • Zeiss AxioObserver for wide-field fluorescence and micro manipulation
  • Zeiss AvioZoom v16 with Apotome 2 for structured illumination
  • microscopes and stereomicroscopes for routine imaging and analysis

Examples of recent work include:

Applicants should submit the following to Dr. David Parichy (dparichy@virginia.edu):

• CV

• contact information for three references

• brief description of interests, experience and career goals

 

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Senior Research Position (Boston, MA)

Posted by , on 27 June 2018

Closing Date: 15 March 2021

The DRSC/TRiP Functional Genomics Resources in the Perrimon group at Harvard Medical School in Boston, MA, USA, is seeking a highly motivated senior-level research technician to join our team. The successful candidate will be responsible for performing molecular biology, cell culture, protein purification, and/or related techniques as part of an overall research program focused on new technology and resource development (e.g. CRISPR technologies, cell-based assays, protein-based studies). The job requires performing independent work that is coordinated with a larger research team. Although most time will be dedicated to hands-on activities, duties will also include quality assessment and data entry.

We provide many opportunities for continued learning in a dynamic work environment. The successful candidate will have access to training on our automated equipment and have the option to attend weekly lab meetings, department seminars, and so on.

You can learn more about our collaborative team, community-focused efforts, and leading-edge research using Drosophila and Drosophila cultured cells at the websites below. Follow the link to the Perrimon lab web page to view details and apply for the job.
https://fgr.hms.harvard.edu/
https://perrimon.med.harvard.edu/

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Blastoid: the backstory of the formation of blastocyst-like structure solely from stem cells.

Posted by , on 27 June 2018

In our recently published paper1, we showed that mouse stem cells self-organize into blastocyst-like structures, that we termed blastoids. Because blastoids can be generated in large numbers, can be finely manipulated, and implant in utero, they are a powerful tool to investigate the principles of pre- and post-implantation development. Here is the backstory of our discovery and why we think it is important for science and medicine.

Nature | doi:10.1038/s41586-018-0051-0

 

The early mammalian embryo, known as the blastocyst, combines the simple and esthetic design of an outer cyst (the future placenta) englobing an inner cell cluster (the future embryo) with the powerful potential to form the whole organism (both embryonic and extra-embryonic). But blastocysts are also few, small, with a small number of cells (<100), and are difficult to physically and genetically manipulate2. Many of its operating principles therefore remain unknown.

 

Stem cell-based embryology.
Stem cells have the intrinsic capacity to self-organize in vitro, as shown with post-implantation models known as gastruloids3-5 or with organoids. Here, we created the first version of a pre-implantation blastocyst model by promoting the self-organization of mouse embryonic (ESCs) and trophoblast stem cells (TSCs).

The reductionist approach of forming entities ‘from the bottom up’ allows to modulate and reveal previously unnoticed principles of development. In addition, blastoids have technical advantages as compared to blastocysts. Large number of genetically similar structures can be generated, thus opening possibilities for high-throughput screens and for biochemistry-based assays. Also, sub-populations can be rapidly and efficiently fine-tuned. For instance, the trophoblast and embryonic compartments can be physically and genetically modified independently one from another (e.g. dosing/mixing of different genotypes)1, 2.

Altogether, blastoids are models to study the genetically-encoded principles of self-organization, and to generate novel hypotheses on development, which complement the classical ‘top-down’ approach (e.g. observing genetically modified blastocysts). In the future, blastoids, which comprise the cell types necessary to form the whole organism and which can be transferred in utero, might form a full organism solely from stem cells.

 

Pulling forces to initiate the project.

As an undergraduate in France, I studied a range of physics and engineering-related topics such as polymer physics and fluid dynamics, which are statistical and modelling sciences. However, what grasped my imagination was the experimental concept of tissue engineering as proposed by Linda Griffith and Robert Langer6. I applied it during my PhD (2006-2010) by studying the self-organization of pre-vascularized engineered tissues (Prof. Clemens van Blitterswijk, University of Twente, The Netherlands). At the end of it, Clemens van Blitterswijk gave me the freedom to work independently, and to hire two PhD students. A great opportunity to develop new ideas, and at an interesting time: the laboratories of Hans Clevers (Hubrecht Institute) and of Yoshiki Sasai (CDB RIKEN) had just shown that stem cells can form organoids. These discoveries perfectly balanced self-organization and developmental biology.

As I wanted to model the embryo, the blastocyst appeared as a lucid choice in light of the available ESCs and TSCs types. The necessity to gain knowledge on blastocysts stem cells led me to ask Niels Geijsen to become a guest at the Hubrecht Institute for developmental biology and stem cell research (Utrecht, The Netherlands). I obtained an additional grant (“Modulating trophectoderm pluripotency and placental development in artificial blastocysts”, ZonMw project number 116005008), hired two great PhD students Erik Vrij (2011) and Javier Frias Aldeguer (2012), and started pipetting ESCs and TSCs, while finalizing the publication of my PhDs’ papers (2012)7,8. Erik focused on developing high-content imaging to quantify the phenotypes, while Javier took a deep dive into the relatively unexplored biology of TSCs.

It took us years of intense, stressful and risky work, and we had to run less uncertain projects to maintain a reasonable output, but the team progressively gained expertise and developed efficient, robust experimental pipelines.

 

How did it come to work?

Self-organization relies on setting up the right boundary conditions to trigger the process. Once initiated, the stem cells remember where they come from, and unleash their intrinsic potential. Two elements were key in creating these initial conditions, which also apply to other self-organizing biological systems. First, a fine control over the confinement of minute, precise number and ratios of ESCs and TSCs. We achieved this using a hydrogel microwell array that I designed and fabricated during my PhD8. Second, the possibility to screen for combinations of molecules to trigger the process. We did this by combining transcriptomic databases with the knowledge on signaling pathways previously found by many blastocyst and stem cells labs. The list is long here but we were definitely standing on the shoulders of Janet Rossant, Jennifer Nichols, Austin Smith.

Our approach suggested new mechanisms. For instance, many Wnt ligands are known to be produced by the blastocyst cells (e.g. Wnt7b and Wnt6 by the trophoblasts9) but their functions are not known (knock-out mice are not informative10, possibly due to the plasticity and redundancy of pathways). I clearly remember when stimulating the cells with Wnt activators (Wnt3a, CHIR99021) and looking at the plates 48 hours later. This triggered the cavitation of TSCs, resulting in gorgeous trophectoderm-like cysts. We thought “phenotypically, it is becoming really good and efficient. We might get somewhere, but are we looking at something really occurring in the trophectoderm?” At the moment, no-one knows but blastoids allow to generate such hypotheses that couldn’t be tested until now.

Once the initial conditions are gathered, including a cocktail of six molecules, the stem cells spontaneously organize within 65 hours. The process is rapid and efficient: about 70% of the microwells that include an adequate number of stem cells form a blastoid (see our definition of a blastoid1).

 

Making discoveries at the single cell level.

In the meantime, the Hubrecht Institute changed director. Hans Clevers stepped down to focus on national-level activities, and Alexander van Oudenaarden came back from MIT to take the job. He set up a large lab focusing on single cell technologies, which attracted our attention as a powerful way to reveal the gene expression patterns underlying the self-organization of blastoids.

We undertook a series of experiments with Jean-Charles Boisset, postdoc in Alexander’s lab. The initial bulk sequencing run comparing blastoids and blastocysts was probably the second convincing moments that made us feel that we were going in the right direction. With his natural phlegm, Jean-Charles pushed the button on his computer to generate the distance map and simply said “it seems to work”, as a heat-map came out on his screen. The transcriptome of blastoid cells was shifting toward the one of a blastocyst, and we had the green light to depict the phenomena at the single cell level. By physically decoupling the compartments (ESCs or TSCs alone versus blastoid cells) and comparing the single cells with the ones from blastocysts, we pinpointed at changes and at the role of the communication between the TSCs and ESCs.

The ESCs clearly played an inductive role: maintaining trophoblast proliferation, inducing morphogenesis, gate-keeping the progression of trophoblast differentiation, and altogether preserving the potential for the trophectoderm to implant in utero. We established a long list of embryonic inductions that guide trophectoderm development (e.g. metabolic, Jak/STAT, Wnt, SMAD, and Hippo pathways), all of which were interesting mechanisms probably contributing to the formation and implantation of the blastocyst.

 

Reviewing, forever.

It took us two years to convince the reviewers that the blastoid system was modeling relevant aspects of blastocyst development. We were asked, among many other experiments, to obtain phenotypes upon generation of KO within ESCs and within TSCs. We replaced the ESCs by other cell types to prove the specificity of the embryonic inductions, by the factor that they secrete as well (BMP4, Nodal), and ran high-throughput phenotypic screens to quantify the morphogenetic and functional impact of embryonic inducers.

The in-utero transfer assay was probably crucial to tip the balance as, for the first time, stem cells alone formed a full entity that could be transferred and tested in utero. We did not form a bona fide embryo in utero but blastoids implanted and induced the expression of Aldh3a1 in the deciduae, which is thought to be a specific response to blastocyst implantation (as compared to polymer beads)11. Blastoid cells proliferated, elongated and generated multiple relevant cell types, including giant trophoblasts that hooked up with the mother’s blood system.

Finally, the paper was released in May 2018, and gained attention from the media. I interviewed for the BBC, BBC World News (live!), CNN or Fortune, and more than 100 press articles were released worldwide. We managed to restrain the unfounded fantasy for unethical human reproduction approaches, and to focus on the opportunities to research in the lab the fundamental principles of development or the minor flaws that can occur at the start of pregnancy. These flaws can prevent the conceptus to implant or can contribute to sub-optimal pregnancies (e.g. sub-optimal placenta development) affecting the appearance of chronic diseases during adult life. The biology community also efficiently relayed realistic and positive potential impacts. As free-electrons in stem cell biology and embryology, it has been gratifying to see the positive and encouraging reactions of valued scientists12-14. A Tweet worth years of research15.

 

Imagining the future of blastoids.

Along the way, curiosity led me to discuss with clinicians including human geneticists, epidemiologists, or IVF clinicians. We started to think of realistic possibilities to use blastoids to research in the lab the problems of infertility, contraception, or early pregnancy failure12. These are extensive societal problems: On one side, humans, which are poorly efficient at procreating, currently delay more and more their pregnancy, which leads to a drop of fertility16. On the other side, family planning and contraception remains a major global health problem as depicted by WHO17 and the Bill & Melinda Gates foundation18.

Overall, women must be able to better plan their pregnancy without decreasing their chance of having a child. Family planning is a huge lever to secure women’s autonomy and well-being, while supporting the health and development of communities. There is a long way in front of us but we are thrilled to see that we might be able to reveal new principles in embryology and, along with clinicians, tackle global health problems.

 

We are currently recruiting Postdocs and PhD students. Feel free to contact me for more information. www.nicolasrivron.org

 

[1] Rivron NC [corresponding author], Frias-Aldeguer J, Vrij EJ, Boisset JC, Korving J, Vivié J, Truckenmüller RK, van Oudenaarden A, van Blitterswijk CA †, Geijsen N † [† equal contribution]. Blastocyst-like structures generated solely from stem cells. Nature. 2018. 557, pages106–111 (2018). doi:10.1038/s41586-018-0051-0.

[2] Rivron NC. Formation of blastoids from mouse embryonic and trophoblast stem cells. Protocol Exchange (2018) doi:10.1038/protex.2018.051

[3] van den Brink SC, Baillie-Johnson P, Balayo T, Hadjantonakis AK, Nowotschin S, Turner DA, Martinez Arias A. Symmetry breaking, germ layer specification and axial organisation in aggregates of mouse embryonic stem cells. Development. 2014 Nov;141(22):4231-42. doi: 10.1242/dev.113001.

[4] Harrison SE, Sozen B, Christodoulou N, Kyprianou C, Zernicka-Goetz M. Assembly of embryonic and extraembryonic stem cells to mimic embryogenesis in vitro. Science. 2017 Apr 14;356(6334). pii: eaal1810. doi: 10.1126/science.aal1810.

[5] Warmflash A, Sorre B, Etoc F, Siggia ED, Brivanlou AH. A method to recapitulate early embryonic spatial patterning in human embryonic stem cells. Nat Methods. 2014 Aug;11(8):847-54. doi: 10.1038/nmeth.3016.

[6] http://news.mit.edu/2012/engineering-health-tissue-engineering-growing-organs-1214

[7] Rivron NC, Raiss CC, Liu J, Nandakumar A, Sticht C, Gretz N, Truckenmüller R,  Rouwkema J, van Blitterswijk CA. Sonic Hedgehog-activated engineered blood vessels enhance bone tissue formation. Proc Natl Acad Sci U S A. 2012 Mar 20;109(12):4413-8. doi: 10.1073/pnas.1117627109.

[8] Rivron NC, Vrij EJ, Rouwkema J, Le Gac S, van den Berg A, Truckenmüller RK, van Blitterswijk CA. Tissue deformation spatially modulates VEGF signaling and angiogenesis. Proc Natl Acad Sci U S A. 2012 May 1;109(18):6886-91. doi: 10.1073/pnas.1201626109.

[9] Kemp C, Willems E, Abdo S, Lambiv L, Leyns L. Expression of all Wnt genes and their secreted antagonists during mouse blastocyst and postimplantation development. Dev Dyn. 2005 Jul;233(3):1064-75.

[10] van Amerongen R, Berns A. Knockout mouse models to study Wnt signal transduction. Trends Genet. 2006 Dec;22(12):678-89.

[11] McConaha, M. E., Eckstrum, K., An, J., Steinle, J. J. & Bany, B. M. Microarray assessment of the influence of the conceptus on gene expression in the mouse uterus during decidualization. Reproduction 141, 511–527 (2011).

[12] Rossant J, Tam PPL. Exploring early human embryo development. Science. 2018 Jun 8;360(6393):1075-1076. doi: 10.1126/science.aas9302.

[13] https://f1000.com/prime/733152526#eval793546032

[14] http://www.sciencemediacentre.org/expert-reaction-to-study-reporting-the-development-of-embryo-like-structures-derived-from-mouse-stem-cells/

[15] https://twitter.com/AMartinezArias/status/991729701130563584

[16] https://www.nytimes.com/2018/05/17/us/fertility-rate-decline-united-states.html

[17] http://www.who.int/news-room/fact-sheets/detail/family-planning-contraception

[18] https://www.gatesfoundation.org/What-We-Do/Global-Development/Family-Planning

 

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Editorial: Advocating developmental biology

Posted by , on 27 June 2018

This editorial by Aidan MaartensAndreas ProkopKatherine Brown and Olivier Pourquié originally appeared in Development


 

Developmental biology is a discipline with a long and rich history, a vibrant and diverse present, and a future of tremendous potential. The field has had enormous impact beyond its own boundaries, for example providing many key concepts for medical research and laying the foundations for advances in the stem-cell and tissue-engineering fields. Technological advances are bringing new solutions to problems that have preoccupied the field for decades, including the potential to analyse our own (human) development. We stand poised on the brink of a deeper understanding not only of development, but increasingly also of regeneration and ageing.

However, while we and others (St Johnston, 2015; Gilbert, 2017) would argue that the field is in a strong position, there is also reason for concern. At conference poster sessions and coffee breaks, departmental happy hours and water coolers, conversations inevitably turn to questions of funding and the future – the next grant, the next position, the next budget. Young researchers look to an uncertain future, and reasonably consider their place in it. Some countries are hit harder than others, and some researchers are more worried than others, notably those conducting basic research without an immediate translational impact. Of course, developmental biologists are not unique in feeling these concerns, but this does not lessen the stark contrast between the promise of the discipline and the threats felt against it. It is therefore vital to consider what individuals and organisations can do to advocate for the continuing importance of developmental biology. This editorial discusses some of our efforts in this regard, and announces a new article series that we hope will provide a useful advocacy resource for the field.

Over the years, A.P. [Communications Officer of the British Society for Developmental Biology (BSDB, www.bsdb.org/)] has engaged in numerous long-term outreach and advocacy initiatives, and highlighted the importance of developmental biology (www.openaccessgovernment.org/developmental-biology-important/41386/). He argues that we should be ready to stand up for our field whenever there is an opportunity to engage with audiences, including the wider public, students, clinicians, journalists, funding agencies and policy makers. We need effective ways to do this, and our engagement will become more powerful if we collaborate and share our strategies and resources (see Illingworth and Prokop, 2017 and references therein). One concrete suggestion from A.P. is to prepare elevator pitches that convincingly explain the importance of your research; thenode.biologists.com/advocacy/outreach/provides a concise rationale and selection of ideas for such pitches, as well as numerous references for individuals to further strengthen their case. So, reader, do you know your elevator pitch, and are you ready to engage?

The BSDB’s advocacy initiative is just one example of how organisations can help advocate developmental biology. The Society for Developmental Biology in the USA (www.sdbonline.org/) has undertaken numerous education and outreach activities, and with the BSDB and other societies is part of the global umbrella organisation the International Society for Developmental Biology (www.developmental-biology.org/). In the field of stem cell biology, the International Society for Stem Cell Research (www.isscr.org/about-isscr) provides a platform for advocacy, education about the latest stem cell advances. As well as facilitating science communication activities, such professional societies, along with funding bodies and academic institutions, can also do the essential (but perhaps less well documented) work of actively liaising with policy makers to achieve the necessary recognition of the importance of our discipline.

But what can journals do? Development has long prided itself as a community journal, and seeks to help researchers in many ways aside from publications, for instance through our travelling fellowships and meeting grants supported by our not-for-profit publisher The Company of Biologists. Through social media, we promote inspiring new research to specialists and non-specialists alike. We also host a community blog, the Node (thenode.biologists.com), which serves as a space for developmental biologists to share information and ideas, and could, in principle, provide a platform for individuals and societies to cooperate and coordinate their advocacy efforts. To help facilitate this, Sarah Morson (who joined us in 2017 for a three-month internship) updated the Node’s Resources page (available at thenode.biologists.com/resources). This resource incorporates the collections of links previously managed by A.P. on the BSDB website, covering the areas of advocacy, outreach and education, as well as sections dedicated to audio-visual and research tools. We hope these pages will be a dynamically evolving resource and encourage suggestions from the community for useful additions.

With its wide readership, Development can act as a mouthpiece to advocate our discipline; that we should get more involved in such efforts was a strong theme running through the feedback we received during our recent community consultation. Following discussions among ourselves and with the journal’s editorial group, we are now pleased to announce a series of articles that aim to advocate the wider importance of developmental biology.

The first set of articles will set out to answer the question ‘What has developmental biology ever done for us?’ with a series of case studies linking particular discoveries in developmental biology to their wider scientific and societal impact. Most obviously, this will involve examples of how basic knowledge gleaned from model organisms has led to medical applications, but we also want to emphasise how the field has advanced our general understanding of how life works, contributing to knowledge and education as social values in their own right (Rull, 2014). With this set of articles, we aim not only to celebrate the prestigious history of our field, but also to provide concrete examples showing why we need to continue to do basic developmental biology research.

To complement this historical angle, the second set of articles will look forward and ask ‘What are the big open questions in the field?’ We want to explore the fundamental unanswered questions in developmental biology and propose how we might start addressing them. The aim is not to wallow in the mystery of these questions, but rather to argue that, particularly with the breakneck speed of development of new tools, they are increasingly tangible. As well as providing signposts for the field’s future, we hope that these articles will convince prospective students that there has never been a more exciting time to get involved in developmental biology – and, perhaps, provide inspiration in choosing their particular field of research.

By providing a rationale for why we do developmental biology and where it is taking us, we hope that these articles will help to advocate our discipline, providing a useful resource for developmental biology educators and advocates, and also helping current researchers to develop their elevator pitches. Our first articles appear in this issue: Katrin Wiese, Roel Nusse and Renée van Amerongen survey the history of the Wnt pathway through multiple model organisms to its influence on the cancer and stem cell fields (Wiese et al., 2018), while Miki Ebisuya and James Briscoe provide a perspective on the meaning of time development (Ebisuya and Briscoe, 2018). We hope you will enjoy this collection of articles, and we welcome suggestions for further commissions.

Developmental biology is a vital, fascinating and evolving discipline. As a global community, we can help to safeguard and support our field going forwards, and we encourage you all – through outreach and communication activities, discussions with funders and policy-makers – to get involved.


 

To see the full collection as it grows, go to

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Visualizing data with R/ggplot2 – One more time

Posted by , on 26 June 2018

Experiments are rarely performed in isolation. Usually, several conditions are compared in parallel or sequential experiments. This experimental strategy also applies to time-dependent data, e.g. from timelapse imaging. So, naturally, after I published a ‘walk-through for plotting temporal data using R and ggplot2, I was immediately asked how to plot two (or more) sets of data in the same graph.

To respond to this request, I made a ‘walk-through’ for generating graphs that visualize temporal changes in signal from multiple experimental conditions. This tutorial will also feature some of the ‘tidy tools’ (footnote 1) that are designed to work on tidy data. The final figure, shown below, displays the changes in the activities of three (separately measured) Rho GTPases that occur after stimulating endothelial cells. For more background on the experiments, the reader is referred to the paper by Reinhard et al (2017).

Combining and plotting the data from different conditions

First we will read the experimental data from three different csv files (available here) and put them into three separate dataframes:

> df1 <- read.csv("Fig3_Rac_S1P.csv")
> df2 <- read.csv("Fig3_Rho_S1P.csv")
> df3 <- read.csv("Fig3_Cdc42_S1P.csv")

Each dataframe holds the information from a particular experimental condition. In this example, the dataframes contain the activities of three Rho GTPases, Rac, Rho or Cdc42, measured over time. Each dataframe has data from individual cells organized by columns. The values that were measured are a “Ratio” that is a result of a FRET imaging experiment. The first column is a record of the Time. Let’s see what a typical dataframe looks like:

> head(df1)

returns:

       Time    Cell.1    Cell.2    Cell.3    Cell.4    Cell.5 ...
1 0.0000000 1.0012160 1.0026460 1.0022090 0.9917870 0.9935569 ...
2 0.1666667 0.9994997 0.9928106 0.9997658 0.9975348 1.0018910 ...
3 0.3333333 0.9908362 0.9964057 0.9905094 0.9946743 0.9961497 ...
4 0.5000000 0.9991967 0.9972504 0.9972806 1.0074250 1.0060510 ...
5 0.6666667 1.0093450 1.0109910 1.0103590 1.0084080 1.0022130 ...
6 0.8333333 0.9941078 0.9940830 0.9990720 1.0181230 1.0110220 ...

To keep track of which data or values belong to which experimental condition (Rac, Rho or Cdc42) we assign a unique identifier to each of the dataframes. To achieve this, we add another column, named “id”, that identifies the condition:

> df1$id <- "Rac"
> df2$id <- "Rho"
> df3$id <- "Cdc42"

The next step is to merge the datasets into one dataframe. The function bind_rows() from the ‘dplyr’ package will do this (to learn how to load the package see footnote 1):

>df_merged <- bind_rows(df1,df2,df3)

Columns with identical names will be merged. For instance, there will be one column with “Time” and another column named “id” that indicates which experiment was performed. The remaining columns list the values that were obtained during the measurements for each of the cells.

The dataframes  from the different conditions have data from different numbers of cells. Therefore, there will be columns that are not completely filled with values. For instance, the dataframe df1 (Rac) has data from 32 cells, while df2 (Rho) has only data from 12 cells. As a result, the df_merged dataframe has a column “Cell.32” that has only data from the Rac experiment and not for Rho and Cdc42. These empty fields (or “missing values”) will be replaced in the dataframe by “NA”.

To do all of the above at once for all csv files that are present in a defined working directory, this R script can be used.

The merged dataframe, df_merged, is still in a wide, spreadsheet format. To convert it into a tidy format (as has been detailled here) we use the function gather() from ‘tidyr’. Since the “Time” and “id” column do not need to be changed, they are excluded from the gathering operation by the preceding minus sign:

>df_tidy <- gather(df_merged, Cell, Ratio, -id,-Time)

To check the result use:

> head(df_tidy)

which returns:

       Time  id   Cell     Ratio
1 0.0000000 Rac Cell.1 1.0012160
2 0.1666667 Rac Cell.1 0.9994997
3 0.3333333 Rac Cell.1 0.9908362
4 0.5000000 Rac Cell.1 0.9991967
5 0.6666667 Rac Cell.1 1.0093450
6 0.8333333 Rac Cell.1 0.9941078

Previously, we used a similar tidy dataframe to draw the lines from individual cells and color those according to the column “Cell”:

>ggplot(df_tidy, aes(x=Time, y=Ratio)) + geom_line(aes(color=Cell))

This results in:

This graph looks like a mess. The reason is that in the column “Cell”, the factor Cell.1 can be linked to the Rho, Rac and Cdc42 dataset.

To solve this, we need to define another column that has a unique identifier for each cell from every condition. One way to achieve this, is by combining the string in the “id” column with the string in the “Cell” column by using unite(). The new, combined string (connected with an underscore) is placed into a new column named “unique_id”:

>df_tidy <- unite(df_tidy, unique_id, c(id, Cell), sep="_", remove = FALSE)

Now, each sample can be grouped according to the “unique_id” to draw the line correctly:

>ggplot(df_tidy, aes(x=Time, y=Ratio)) +
  geom_line(aes(color=Cell, group=unique_id))

To obtain a plot that shows all the individual traces colored according to the experimental condition we can use:

>ggplot(df_tidy, aes(x=Time, y=Ratio)) + geom_line(aes(color=id, group=unique_id))

 

 

Generating a graph with data summaries

The next thing you may want to do is to calculate statistics. In the previous post it was explained how to calculate the mean value for each time point by using base R code.

Here, the same calculation will not give a meaningful result, since each time point has data from the three different conditions. To calculate the mean for each condition (defined by “id”) for each of the time points I will use the function group_by(). This function is part of the “tidyverse”, a set of tools that work well on tidy data (footnote 2).

Another feature of tidy tools is the “pipe” which is encoded by %>%. It basically moves the result from a function onto the next function. The advantage is that we can concatenate a number of functions that are still understandable by humans because the sequence of events can be read from left to right (footnote 3).

So, to calculate the mean value we first group the data by “Time” and “id”. Next, we calculate the number of points (n) and the average (mean) with the function summarise(). The pipe is used to perform the operations all at once. The result is assigned to a new dataframe, df_tidy_mean:

>df_tidy_mean <- df_tidy %>%
    group_by(Time, id) %>%
    summarise(n = n(), mean=mean(Ratio))

To show what the first rows of the dataframe looks like we use head():

> head(df_tidy_mean)

which yields:

# A tibble: 6 x 4
# Groups:   Time [2]
Time id        n   mean
<dbl> <chr> <int>  <dbl>
1 0     Cdc42    32 NA
2 0     Rac      32  0.998
3 0     Rho      32 NA
4 0.167 Cdc42    32 NA
5 0.167 Rac      32  0.999
6 0.167 Rho      32 NA

The data structure of the result is a “tibble“, which is a next-generation dataframe. In this walk-through, we use tibbles in the same way as dataframes and do not make a distinction.

The content of df_tidy_mean is not what we want, since each row has n=32 observations and the mean of de conditions “Cdc42” and “Rho” returns “NA”. We know that n=32 for Rho cannot be right, since there are only 12 cells cells measured for Rho. The n=32 for each of the conditions is due to the “NA” values in the columns that are counted as well. This also explains why we do not see a proper mean for “Cdc42” and “Rho”. To address this issue, we need to get rid of the missing values in the “Ratio” column. We can do this by adding another function to the previously used command that uses the filter() function to only select “Ratio” values that are not “NA”:

> df_tidy_mean <- df_tidy %>%
    filter(!is.na(Ratio)) %>%
    group_by(Time, id) %>%
    summarise(n = n(), mean=mean(Ratio))

Let’s see if that worked out the way we want:

> head(df_tidy_mean)
# A tibble: 6 x 4
# Groups:   Time [2]
Time id        n  mean
<dbl> <chr> <int> <dbl>
1 0     Cdc42    19 1.00
2 0     Rac      32 0.998
3 0     Rho      12 0.998
4 0.167 Cdc42    19 0.995
5 0.167 Rac      32 0.999
6 0.167 Rho      12 1.00

This gives the expected result. The condition “Rho” has n=12 and the condition “Rac” has n=32. These number of samples match with the data (csv files) that were supplied as input. We can continue to add some more useful statistics to the dataframe:

>df_tidy_mean <- df_tidy %>%
    filter(!is.na(Ratio)) %>%
    group_by(Time, id) %>%
    summarise(n = n(),
           mean = mean(Ratio),
         median = median(Ratio),
             sd = sd(Ratio))

Finally, we will use the function mutate() to calculate the inferential statistics, i.e. the standard error of the mean and the 95% confidence interval of the mean:

>df_tidy_mean <- df_tidy %>%
    filter(!is.na(Ratio)) %>%
    group_by(Time, id) %>%
    summarise(n = n(),
           mean = mean(Ratio),
         median = median(Ratio),
             sd = sd(Ratio)) %>%
    mutate(sem = sd / sqrt(n - 1),
      CI_lower = mean + qt((1-0.95)/2, n - 1) * sem,
      CI_upper = mean - qt((1-0.95)/2, n - 1) * sem)

Now, the df_tidy_mean dataframe has all the summary data that we need, for each condition (“id”) and for each time point.

> head(df_tidy_mean)
# A tibble: 6 x 9
# Groups:   Time [2]
Time id        n  mean median      sd      sem CI_lower CI_upper
<dbl> <chr> <int> <dbl>  <dbl>   <dbl>    <dbl>    <dbl>    <dbl>
1 0     Cdc42    19 1.00   1.00  0.00756 0.00178     0.999    1.01
2 0     Rac      32 0.998  0.998 0.00550 0.000988    0.996    1.000
3 0     Rho      12 0.998  0.997 0.00477 0.00144     0.995    1.00
4 0.167 Cdc42    19 0.995  0.997 0.00565 0.00133     0.993    0.998
5 0.167 Rac      32 0.999  0.999 0.00684 0.00123     0.997    1.00
6 0.167 Rho      12 1.00   1.00  0.00615 0.00186     0.999    1.01

That’s all we need to make a graphical summary of the data:

>ggplot(df_tidy_mean, aes(x=Time, y=mean, color = id)) +
  geom_line(aes(x=Time, y=mean, color=id)) +
  geom_ribbon(aes(ymin=CI_lower,ymax=CI_upper,fill=id),color="grey70",alpha=0.4)

 

Improving the presentation and annotation of the graph

Further styling of the figure can be done. Below, we get rid of the default ggplot2 theme, change the scale of the x-axis and add a vertical line at t=1.75 to indicate when the stimulation took place. This will generate the final figure that was shown above:

>ggplot(df_tidy_mean, aes(x=Time, y=mean, color = id))+
  geom_line(aes(x=Time, y=mean, color=id))+
  geom_ribbon(aes(ymin=CI_lower,ymax=CI_upper,fill=id),color="grey",alpha=0.4)+
  theme_light(base_size = 16) + xlim(0,10) + geom_vline(xintercept = 1.75)

Color

User defined colors can be added to the lines and ribbons by using the functions scale_color_manual() and scale_fill_manual() respectively. The list of color values can consist of colornames or hexadecimal RGB code:

>color_list <- c("#EE6677", "#228833", "#4477AA")

Or:

>color_list <- c("darkgoldenrod", "limegreen", "turquoise2")

When the colors are defined, they can be used for plotting:

>ggplot(df_tidy_mean, aes(x=Time, y=mean, color = id))+
  geom_line(aes(x=Time, y=mean, color=id), size=1)+
  geom_ribbon(aes(ymin=CI_lower,ymax=CI_upper,fill=id),color="grey",alpha=0.4) +
  theme_light(base_size = 16) + xlim(0,10) + geom_vline(xintercept = 1.75) +
  scale_fill_manual(values=color_list) + scale_color_manual(values=color_list)

 

Side-by-side

Thus far, we made a single graph for all the data. To see the data from the different conditions (“id”) in separate plots, we add “facet_grid(.~id)”:

>ggplot(df_tidy_mean, aes(x=Time, y=mean, color = id))+
  geom_line(aes(x=Time, y=mean, color=id), size=1)+
  geom_ribbon(aes(ymin=CI_lower,ymax=CI_upper,fill=id),color="grey",alpha=0.4) +  theme_light(base_size = 16) + xlim(0,10) + geom_vline(xintercept = 1.75) +
  scale_fill_manual(values=color_list) + scale_color_manual(values=color_list) +
  facet_grid(.~id)

The result is three plots next to each other (in a horizontal format). However, in this case it probably makes more sense to arrange the plot in a single column. This is achieved by adjusting the order between the brackets: facet_grid(id~.). Since a legend seems redundant, it is removed in the following example:

>ggplot(df_tidy_mean, aes(x=Time, y=mean, color = id))+
  geom_line(aes(x=Time, y=mean, color=id), size=1)+
  geom_ribbon(aes(ymin=CI_lower,ymax=CI_upper,fill=id),color="grey",alpha=0.4) +  theme_light(base_size = 16) + xlim(0,10) + geom_vline(xintercept = 1.75) +
  scale_fill_manual(values=color_list) + scale_color_manual(values=color_list) +
  facet_grid(id~.)+ theme(legend.position="none")

 

Final words

The use of transparent layers for the different conditions allows to combine the data in a single graph. The resulting plot provides a clear view of the relation between the activities. Alternatively, the plots can be readily shown as a column in which the time axes are aligned (similar to the original figure 3 from Reinhard et al., 2017). I hope that providing this ‘walk-through’ that shows how to build a graph from multiple datasets will be a good starting point to generate plots of your own data with R/ggplot2.

Acknowledgments: Thanks to Eike Mahlandt and Max Grönloh for testing and debugging the code.

 

Footnotes

Footnote 1: The R packages that are necessary for this walk-through are: ‘dplyr’, ‘tidyr’, ‘ggplot2’ and ‘magrittr’. To load a package (after downloading and installing it) use require(), for example:

>require('dplyr')

The aforementioned packages are part of the ‘tidyverse’ and can be installed all at once.

Footnote 2: I usually mix basic R functions with functions from tidytools. This is mostly for ‘historical’ reasons; I first learned a bit of base R before I learned about the tidyverse. In my experience, some of the base R code is easier to remember and therefore my default code is base R. On the other hand, the tidy tools allow to string a number of functions together (using the pipe; %>%) which condenses the code, while maintaining readability.

Footnote 3: There are many places where the application of tidy tools is explained in more detail, for example here and here.

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BBSRC-funded postdoc position – neuronal ageing

Posted by , on 26 June 2018

Closing Date: 15 March 2021

BBSRC funded postdoc position in the laboratory of Natalia Sanchez-Soriano (https://sanchezlab.wordpress.com), to study the cell biology of neuronal ageing and the underlying mechanisms.  The aim of the project is to understand the harmful changes that neurons undergo at the subcellular level during ageing, and unravel the cascade of events that cause them, with a focus on intracellular degradation systems and the upstream regulatory pathways. Ideally applicants should be trained in neuro- and/or in vivo cell biology, and imaging, and have experience with Drosophila.


The post is available from 1st of September 2018 until 31st of August 2021.
For full details and to apply online, please visit: https://recruit.liverpool.ac.uk
Job Ref: 009401, closing Date: 28 June 2018.

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Postdoctoral fellowship in Applied Developmental Immunology

Posted by , on 22 June 2018

Closing Date: 15 March 2021

The Maehr lab (http://maehrlab.net/) is looking for a team member in the area of developmental immunology and human pluripotent stem cell-based disease models.

Envisioned projects will utilize pluripotent stem cell differentiation approaches, together with assays of thymus and T cell development, to decipher the molecular underpinnings of human immune syndromes such as thymus dysfunction-based autoimmunity and immunodeficiency. In addition to stem cell differentiation and immunological assays, the team will apply technologies such as functional genomins (e.g. CRIPSR) and single cell RNA-sequencing as well as computational approaches.

Candidates should possess a Ph.D. and have a strong background in immunology and/or developmental biology. Experience with immunological assays, stem cell differentiation, bioengineering and/or next generation sequencing assays is desirable. Excellent communication, writing, and collaboration skills are essential.

Interested candidates should email a cover letter, CV, and 3 references to Dr. René Maehr, Associate Professor of Molecular Medicine, UMass Medical School (rene.maehr@umassmed.edu).

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Unraveling tissue interactions coordinating neuromuscular morphogenesis: a journey through serendipity

Posted by , on 20 June 2018

An emerging trend in developmental biology focuses on the role of cell adhesion in modulating tissue morphogenesis. Spectacular advances have shed light on how modulation of adhesion between equivalent cells orchestrates the acquisition of forms. However, how interactions at complex interfaces between distinct neighboring cell types influence tissue growth remains to be elucidated. In a recent paper, staring mostly at a flat muscle that spreads subcutaneously and its partner motor neurons, I investigated how neuromuscular morphogenesis involves complementary activities of the Fat1 Cadherin in three interdependent cell types – myogenic cells, motor neurons, and the connective tissue through which muscles spreads.

 

Tissue-specific activities of the Fat1 Cadherin cooperate to control neuromuscular morphogenesis Françoise Helmbacher, PLoS Biology, May 16, 2018 | doi: 10.1371/journal.pbio.2004734 | PMID : 29768404

 

The story started as I was transitioning from being a post-doc in the lab of Rüdiger Klein in Munich, to establishing my own group in Marseilles. At the time, I was interested in events of communication between motor neuron (MN) populations, whereby a pool of MNs influences the specification of neighbor MNs. I was looking for molecules mediating such function, and Fat1 originally came up as one of my candidate genes. Although Fat1 met many of my criteria, it was initially considered a low priority candidate for the specific question I was asking (which I won’t discuss here). However, there were also good reasons for deciding to work on Fat1, aside from the main project of my freshly established lab: (1) Fat1 was expressed in my favorite MN population (Figure 1), a group of neurons recognizable by the transcription factor they expressed, which innervates a fascinating muscle, the Cutaneous Maximus (CM), (2) there were mouse mutants available (which by chance happened to be already in my former lab), and (3) through pioneer experiments with these mutants, I had gathered preliminary data indicating that Fat1 disruption resulted in defective specification of the CM motor pool and in defective axonal arborization within the target muscle – the phenotype I was expecting.

 

Whatever the underlying gene function, starting a project with a robust in vivo phenotype guaranteed that an interesting story would emerge from it, even though this was not my main project. For this reason, I kept this as my own side project. I had no hypothesis, but it felt like starting a classical forward genetics approach, with the advantage that I already knew the gene identity. This is also when the field of planar cell polarity (PCP) started blooming, mostly through work in Drosophila. The Drosophila Fat Cadherin, initially regarded as a putative tumor suppressor, had been attributed the mysterious function of propagating information on cell polarity within the plane of epithelia across long distances. I was fascinated by these studies, and curious to ask if MNs could propagate some polarity information to their neighbors too. Well, the story took a completely different path!

 

The two first key results that I had obtained were that 1) axons of the motor neuron pool innervating my favorite muscle, the CM, were clearly shorter in Fat1-deficient embryos, when all other nerves appeared unaffected; and 2) expression of one of the markers specifically characterizing this MN population was drastically reduced. Given the fact that at that time, I had selected Fat1 because of its selective expression in subsets of MNs encompassing the CM-innervating pool (marked by Etv4 expression, Figure 1B), these data were compatible with the simplistic and straight forward hypothesis of a cell autonomous activity of Fat1 in MNs. Thus, it looked as if this project would be devoted to finding out whether specification or axonal growth was affected first, and to tackle the mechanism, keeping PCP principles in mind, possibly using a combination of genetics, in vitro, and in vitro approaches.

Figure 1: (A) Scheme representing the cervical spinal cord and the spinal nerves, which converge at the base of the limb, from which limb-innervating nerves emerge. The Etv4-expressing MN pool and its axons are highlighted in red. (B) Fat1 expression in the spinal cord (visualized using a Fat1LacZ allele or by in situ hybridization) highlights a pool of MNs expressing the transcription factor Etv4. Induction of Etv4 expression is dependent on a peripheral signal, GDNF, secreted by the target muscle of this motor pool, the Cutaneous Maximus muscle. Peripheral Fat1LacZ expression is also detected within this muscle and in surrounding connective tissue. (A, B) appeared originally as Fig5A,C, whereas (C) appeared as Fig4A, B in Helmbacher 2018.

 

Prior to jumping to the conclusion that Fat1 indeed controlled MN specification and axon patterning cell autonomously, it was essential to determine whether the target muscle, the CM, had developed normally in Fat1 deficient embryos. Any muscle defect might indeed represent a potential cause – or consequence – of these MN phenotypes, or might even indicate an independent phenotype. With my team, we therefore performed the experiment that changed the course of this story. We visualized muscle development by whole mount in situ hybridization to detect MyoD RNA in myogenic progenitors. To my surprise, this experiment turned out not to yield the anticipated negative result. Instead, Fat1-deficient embryos exhibited a series of very robust muscle defects affecting morphogenesis of subsets of muscles in the face and shoulder, phenotypes that we described in our PLoS Genetics paper in 2013 [1].

 

When we first saw these muscle phenotypes, there was no reason a priori to dissociate our findings on the altered development of a selective group of muscles and associated defects in the corresponding motor neuron population. However, as I was reflecting on the possible significance of these results in terms of health and biomedical implications, and as, in these early days, I was also in a phase of heavy grant writing, I wondered whether these phenotypes could possibly correspond to symptoms of a rare human pathology that would not have been assigned yet any genetic culprit. I thus started exploring the human genome to see where the human FAT1 gene was located, and discovered the OMIM database (Online Mendelian Inheritance in Man). It didn’t take long to realize that the 4q35 region where FAT1 was localized was also known for being subject to chromosomal abnormalities associated with a human muscular dystrophy called Facioscapulohumeral dystrophy (FSHD) [2]. I was really amazed to learn that the muscle wasting symptoms in this disease affected a highly regionalized group of muscles, with a topography reminiscent of what we had seen in Fat1-deficient embryos. I was also intrigued by the fact that additional non-muscular symptoms of this disease, auditory and vascular retinal abnormalities, were emerging as hallmarks of defective planar cell polarity.

 

At that time, this was a completely unexpected finding. First, the FAT1 gene was relatively far from the true FSHD locus (where changes in the D4Z4 macrosatellite repeat array were considered to be the primary event linked to FSHD). This distance (~3Mb) was sufficiently large to guarantee unambiguous genetic mapping of a disease locus, but was small enough for a genetic linkage, and for FAT1 regulation to be affected secondarily to chromatin architecture changes occurring as a result of the FSHD abnormalities (although the concept of topologically associated domains (TADs) was not as popular then as it has become). Even though a possible deregulation/contribution of neighboring genes had been considered, FAT1 had largely been ignored or dismissed. Nothing in the literature at that time indicated any likely involvement of Fat1 in muscle biology, whereas Fat1 disruption in mice had instead been reported to result in severe kidney abnormalities [3], the latter not being part of the panoply of FSHD symptoms.

 

This is when I decided to radically change the topic of the lab. I therefore sought to identify potential collaborators who would help approaching the human disease link. I was lucky to turn to Nicolas Levy, who headed a department of medical genetics at La Timone Hospital in Marseilles, and was really enthusiastic about this new idea. With his help, we were later joined by Marc Bartoli, who started a group on translational myology in Marseilles, and by Julie Dumonceaux (now at UCL, London). To make a long story short, this fruitful collaboration has led to date to 3 papers exploring the possible link between FAT1 dysfunction and FSHD [1, 4, 5]. First, my team provided compelling data indicating that disrupting Fat1 in mice caused muscular and non-muscular phenotypes with an FSHD-like topography [1]. Second, we detected lowered FAT1 RNA and protein levels in affected muscles from FSHD patients with classical diagnosis, at fetal [1] and adult [5] stages. Third, we identified FSHD-like patients carrying pathogenic FAT1 variants, in absence of the traditional FSHD-causal abnormalities [1, 4]. These FAT1 variants included single nucleotide variants affecting RNA splicing or amino-acid structure [4], but also copy number variants deleting a putative cis-regulatory element in the FAT1 locus [1]. Such deletions had the potential to cause tissue-specific depletion of FAT1 expression, thus offering a logical explanation for why some phenotypes resulting from Fat1 deficiency in mice were not part of the clinical picture of FSHD symptoms in human patients.

 

Because the FAT1/FSHD link was so unanticipated and provocative, the road to our first publication turned into a long and emotionally agitated roller coaster ride. As a result, this paper was submitted multiple times, and went through several rounds of revisions for several journals, until it proudly landed in PLoS Genetics in 2013 [1]. As in the meantime we had produced mice with a conditional Fat1 allele, we were able to show in this first study that Fat1 deletion in the myogenic lineage reproduced in part the aberrant dispersion of myogenic cells in the limb observed in constitutive knockouts [1]. I already knew however that this only gave a partial account of the complexity of Fat1 functions. I just figured that it was unnecessarily challenging to simultaneously report the identification of a potential new genetic actor in a human pathology (even when only considered a modifier gene) and the complexity of its biology. This deserved to be studied as a developmental question per se, independently of its relevance for this disease, which somehow is still a controversial issue, despite our genetic evidence obtained with the Levy/Bartoli groups.

 

To approach the complexity of the Fat1-driven control of neuromuscular morphogenetic, the new study I have just published [6] gives a central position to one emblematic couple: the Cutaneous Maximus (CM) muscle, and its cognate MN pool. This MN pool can be recognized through its expression of the transcription factor Etv4, which is essential to specify the cell body position and dendritic architecture of these MNs [7]. The CM muscle emerges from the bulk of forelimb levels myogenic progenitors that undergo long range migration. Unlike other limb muscles, the CM muscle progenitors avoid entering the limb bud. Instead, they undergo a sharp caudal turn and start their subcutaneous progression. Throughout this migration phase, CM progenitors secrete Glial-cell-derived-neurotrophic-factor (GDNF), previously known for its neurotrophic activity and for its roles in motor axon guidance and in CM motor pool specification (as it induces Etv4 expression) [8]. Thus, by way of GDNF production, there is an intimate link between muscle progression and partner MN development. Among the muscles vulnerable to Fat1 deficiency, the CM muscle was also the easiest to approach quantitatively, by measuring the orientation of individual progenitor cells in our first study [1], and with simple area measurements in the new study [6]. Such an analysis illustrates how Fat1 disruption robustly interferes with expansion of the area occupied by Gdnf-expressing myogenic progenitors, by differentiating muscle fibers (Figure 2), as well as by axons of the cognate MNs.

Figure 2: Fat1 disruption robustly alters the subcutaneous expansion of the CM muscle. (A, B) Developing muscles are visualized at two successive stages (~E12.5) in control and Fat1-/- embryos owing to a transgene expressing a nuclear LacZ reporter (Mlc3f-2E) in differentiating muscle cells (driven by regulatory regions of the sarcomeric gene Mlc3f). Top images show the whole left flank of the embryo, whereas bottom images show higher magnifications of the area in the squared box, featuring the CM muscle. The area delineated with yellow, red, and white dotted lines represent respectively the area occupied by body wall (BW) muscles, by differentiated CM muscle fibers, and by CM myogenic progenitors (indicative). (C) Plots representing: (Left plot) the evolution of the CM area (red lines) with respect to the BW area; and (right plot) the ratio CM/BW normalized to the median control ratio, for each embryo (controls, blue dots; Fat1-/-, red dots). (A-C) appeared originally as Fig1A-C in Helmbacher 2018.

 

Given my initial selection of Fat1 for its expression in the pool of MNs innervating the CM muscle, this was a region of non-myogenic Fat1 expression of obvious relevance to the phenotype. I was really excited by the possibility that Fat1 expression in the CM pool of MNs might represent a key driver of CM subcutaneous morphogenesis. Quantifications of morphometric parameters confirmed that MN-specific Fat1 ablation interfered with axonal growth and specification of the CM-innervating MNs. In addition, this also non-autonomously slows the subcutaneous progression of CM myogenic progenitors, hence expansion of the GDNF producing domain, without altering the rate of myogenic differentiation. However, this role turned out to be way more modest than what I initially anticipated, with an effect size considerably smaller than the phenotypes of constitutive mutants. This implied that another site of Fat1 activity had to be making a major contribution to neuromuscular morphogenesis.

 

An unanticipated third site of Fat1 expression that turned out to play a major role is the loose connective tissue (CT) that surrounds muscles. During their collective progression, myogenic progenitors and motor axons explore such connective tissues and select their migration trajectory. The CM progenitors in particular encounter an increasing gradient of Fat1 expression in this tissue. Furthermore, Fat1 expression appears fully preserved in mouse mutants lacking migratory muscles (Met or Pax3 mutants, in which myoblast migration is abrogated), indicating that the main site of peripheral Fat1 expression represented non-myogenic cells. To establish the contribution of CT-Fat1 to muscle morphogenesis, Fat1 activity was ablated in the lateral plate derived mesenchyme, in a domain (driven by Prx1-cre) including the limb and flank connective tissues encompassing a large part of the territory through which the CM migrates. This led to severe non-cell autonomous disruption not only of the progression of CM progenitors and subsequent myofiber elongation, but also of motor axon elongation across the same territory, and of acquisition of CM fate characteristics. Similar results were obtained with a Cre line allowing stage-controlled Tamoxifen-inducible Fat1 excision in Pdgfrα-expressing connective tissue at the time of myoblast migration, with a lesser effect size matching the lower excision efficiency. These data identify connective tissue as a tissue type in which Fat1 activity is mandatory for neuromuscular morphogenesis. They imply that Fat1 signaling activity in CT cells is necessary for them to emit proper signals promoting collective myoblast migration, myogenic differentiation, motor axon growth, and MN specification, with the exact mechanisms underlying these cell interactions remaining to be explored.

Figure 3: Fat1 ablation in the mesenchyme is sufficient to robustly delay the subcutaneous expansion of CM progenitors. (A) Myogenic progenitors of the CM muscle are visualized owing to a GdnfLacZ allele (by Salmon Gal staining) in control and Prx1-cre; Fat1Flox/Flox embryos, at E12.5. (B) Plots representing: (left plot) the evolution of the area covered by CM progenitors relative to the trunk length of control and Prx1-cre; Fat1Flox/Flox embryos, and (right plot) the CM/TL ratio normalized to controls, for each embryo (controls, blue dots; Prx1-cre; Fat1Flox/Flox, red dots). (C) Scheme explaining how the domain of Prx1-cre activity (in mesenchymal cells), in which Fat1 activity is deleted, is visualized owing to the cre-dependent R26-YFP reporter (green), in relation to the domain of GdnfLacZ expression (red, in CM progenitors). (D) The relative thickness of the CM is visualized on embryo sections by immunohistochemistry with anti-beta-galactosidase and anti-YFP antibodies, in control, Prx1-cre; R26YFP/+; GdnfLacZ/+ and Fat1Flox/Flox; Prx1-cre; R26YFP/+; GdnfLacZ/+ embryos. (A, B) appeared originally as Fig7C,D, whereas (C,D,E) appeared as part of Fig8 in Helmbacher 2018.

 

In conclusion, looking back on how the story started, the loop has been closed by the identification of gene functions that underlie the phenotype which initially attracted my attention. Although my intuition that this had to be connected to the neuronal expression of this atypical Cadherin was not completely wrong after all, conditional mutagenesis nevertheless revealed that this function was modest in regard of the unexpected predominant activity in a cell type – connective tissue – that I had initially not considered. I have learned many lessons along the way of this unusual story. Above all, I learned not to fall in love with a favorite hypothesis but instead to be open to what the facts tell us: when the results of an experiment are the opposite of what you expect, this pushes you to think outside the box, and challenges you to remain unbiased in conceiving experimental plans and analyzing data. It is now time to switch scale, and start exploring what happens at the connective tissue muscle interface from a cell biologist’s point of view.

 

References

  1. Caruso, N., et al., Deregulation of the protocadherin gene FAT1 alters muscle shapes: implications for the pathogenesis of facioscapulohumeral dystrophy. PLoS Genet, 2013. 9(6): p. e1003550.
  2. DeSimone, A.M., et al., Facioscapulohumeral Muscular Dystrophy. Compr Physiol, 2017. 7(4): p. 1229-1279.
  3. Ciani, L., et al., Mice lacking the giant protocadherin mFAT1 exhibit renal slit junction abnormalities and a partially penetrant cyclopia and anophthalmia phenotype. Mol Cell Biol, 2003. 23(10): p. 3575-82.
  4. Puppo, F., et al., Identification of variants in the 4q35 gene FAT1 in patients with a facioscapulohumeral dystrophy-like phenotype. Hum Mutat, 2015. 36(4): p. 443-53.
  5. Mariot, V., et al., Correlation between low FAT1 expression and early affected muscle in facioscapulohumeral muscular dystrophy. Ann Neurol, 2015. 78(3): p. 387-400.
  6. Helmbacher, F., Tissue-specific activities of the Fat1 cadherin cooperate to control neuromuscular morphogenesis. PLoS Biol, 2018. 16(5): p. e2004734.
  7. Livet, J., et al., ETS gene Pea3 controls the central position and terminal arborization of specific motor neuron pools. Neuron, 2002. 35(5): p. 877-92.
  8. Haase, G., et al., GDNF acts through PEA3 to regulate cell body positioning and muscle innervation of specific motor neuron pools. Neuron, 2002. 35(5): p. 893-905.

 

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Lighting Up the Central Dogma in Development

Posted by , on 19 June 2018

We recently published a manuscript in Cell that describes a method to image transcription factor concentration dynamics in real time, in living embryos, using a nanobody-based protein tag that we call the “LlamaTag.” We were particularly excited about these investigations because this new technology overcomes a major technical obstacle to understanding how gene-expression dynamics are regulated in development, which has held back the field for decades.

 

From Snapshots of Dead Embryos to Movies

Remember when you were first starting to study developmental biology? In one of your lab sections, you probably participated in that classic teaching activity where you watched the cephalic furrow form in Drosophila embryos. Maybe you were one of those students struggling to move embryos with an eyelash glued to a toothpick, and the whole time you were thinking, “I need to hurry up—this thing is gastrulating while I’m messing around here!” There’s a strong contrast between watching development unfold before your eyes and scrutinizing snapshots of FISH data from dead, fixed embryos. This contrast hinders more than budding scientific excitement: how can we truly understand dynamic processes by analyzing static data?

I was particularly struck by this conundrum as a postdoctoral researcher as my mind kept replaying a video of a gastrulating embryo. How could I bridge this gap so that developmental biologists could accurately model—and ultimately construct—dynamic systems as complex as embryos? At the simplest level, we would need to know the input and the output of each genetic circuit as it changes throughout development in order to deduce the logic gates at play. I tackled the output end first: as a postdoc, I developed a technique to quantify transcription in flies at the single-cell level by tagging a gene’s nascent mRNA molecules with fluorescent proteins. How could we accomplish the same feat with the DNA-protein interactions that constitute the input to these genetic circuits? I decided that solving this challenge would be a major driver of the research in my own lab. Jacques Bothma was equally excited about this challenge and decided to join the project as a postdoctoral fellow.

Typically, one would quantify the concentration of a transcription factor by fusing it to a fluorescent protein. However, in flies, worms, fish, and frogs, these fluorescent take more than 40 minutes before they mature and become fluorescent. This delay is actually a major problem: the activators and repressors that drive development often exist for less than 10 minutes before they are degraded. By the time the fluorescent proteins actually became fluorescent, the action they were supposed to report on is already over!

Jacques had a great idea: instead of relying on the synthesis of fluorescent proteins, why not use the localization of already matured fluorescent proteins to report gene expression? Our Cell paper describes the engineering and implementation of this idea with LlamaTags, a technology that enables quantitation of the dynamics of transcription-factor activity without limits from the slow maturation of traditional fluorescent fusion proteins. Instead of these fusions, we employed nanobodies, which are small, highly specific, single-domain antibodies that are raised in llamas (hence LlamaTags). We fused a transcription factor of interest (we started with Hunchback) to a nanobody raised against eGFP, and expressed the construct from the endogenous locus for that transcription factor. Importantly, the embryo was engineered to contain maternally deposited eGFP, which means that eGFP is already mature before the transcription factor of interest is expressed (no waiting around for the fusion to mature!). When the transcription factor-nanobody is translated, it binds cytoplasmic eGFP within seconds, yielding a quantitative increase in nuclear fluorescence when the transcription factor moves to the nucleus to perform its regulatory function. LlamaTags therefore deliver a direct readout of the instantaneous transcription-factor concentration in a given nucleus.

Through a variety of experiments, we showed that LlamaTags serve as specific and faithful reporters of the endogenous concentration dynamics of transcription factors during development, thus capturing the input pertinent to these circuits. So, we finally had the two pieces needed to solve the puzzle of measuring input-output functions. LlamaTags made it possible to measure the input concentration of transcription factors in individual nuclei, while MS2 revealed the transcriptional activity of specific genes to these input levels.

By fusing a LlamaTag to Fushi-Tarazu (Ftz), we obtained measurements of its nuclear concentration during development, from which we extracted the in vivo degradation rate of the construct. We were pleased when our data revealed rapid fluctuations in Ftz concentration—consistent with previous reports of protein bursts that likely arise from stochastic fluctuations in mRNA concentrations. We nailed down this relationship, and uncovered exciting evidence of inter-nuclear communication within the embryo. This coupling could be a major driver of the sharp boundaries that dictate development of the fly embryo. However, our greatest satisfaction came from simultaneously visualizing input transcription-factor activity and output transcription, at the single-cell level, in real time, as stripe 2 of eve was laid down in live embryos:

 

Spatiotemporal evolution of the Kr protein expression pattern (green) and eve stripe 2 mRNA expression pattern (red puncta) during the course of nc14. Video S6 in the paper

 

A New “Microscope” for the Central Dogma in Development

Our recent work establishes a powerful pair of technologies—labeling transcription with MS2 and labeling DNA-protein interactions with LlamaTags—that together constitute a “microscope” for visualizing and interrogating the activity of genetic circuits as they function. We envision that LlamaTags can be applied to quantitatively measure the flow of information along regulatory networks in any multicellular organism that is amenable to transgenic control and live imaging. LlamaTags literally light up the central dogma, in real time, as development unfolds. Importantly, we can do more than map input to output: we can quantitate the dynamics of these connections over time. That’s the difference between trying to create a stop-motion movie where you need a new actor for each frame (a dead embryo), and actually observing development unfold in real time.

 

Physical Biology of Living Embryos

Like many of you, we believe that in order to construct a system, we should first understand it in quantitative detail. We view this as a call for reaching a “predictive understanding” of development, through which we can calculate developmental outcomes from knowledge of the concentrations of input transcription factors and the DNA regulatory sequence. To reach this predictive understanding, we believe that a powerful dialogue is necessary in which theoretical models make predictions that are subsequently tested experimentally, with measurements fed forward into the model to generate a new cycle of experiments. Our work in Cell is a crucial early step toward enabling the biophysical dissection of developmental programs by making it possible to measure the very same input-output functions predicted by our theoretical models.

 

 

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Gene Regulatory Networks for Development

Posted by , on 19 June 2018

Applications are now open for this year’s Gene Regulatory Networks for Development which will be at The Marine Biological Laboratory in Woods Hole, USA from October 14- 27.  The application deadline is July 20.  The course is for graduate students, postdoctoral researchers, and faculty members and it focuses on using experimental data and computational modeling to analyze gene regulatory networks that are key to development in animals and plants. 

This unique course is an intense and always interesting experience and has drawn rave reviews in all of its previous incarnations. Students will meet with experts in the field for an in-depth treatment of experimental and computational approaches to GRN science. Through lectures, highly interactive discussions, and group projects we will explore the GRN concept and how it can be applied to solve developmental mechanisms in various systems and contexts. Topics include structural and functional properties of networks, GRN evolution, cis-regulatory logic, experimental analysis of GRNs, examples of solved GRNs in a variety of developmental contexts, and the computational analysis of network behaviour by continuous and discrete modelling approaches. 

Travel fellowships are available.

For more information about the course, go to www.mbl.edu

The 2018 GRN course faculty:

Scott Barolo, University of Michigan

James Briscoe, The Francis Crick Institute, London

Fernando Casares, Andalusian Center for Developmental Biology, Spain

Ken Cho, University of California, Irvine

Doug Erwin, Smithsonian Institution

Robb Krumlauf, Stowers Institute

Bill Longabaugh, Institute for Systems Biology, Seattle

Lee Niswander, University of Colorado, Denver

Isabelle Peter, Caltech

John Reinitz, University of Chicago

Ellen Rothenberg, Caltech

Trevor Siggers, Boston University

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