The successful candidate will participate in the cardiovascular research project funded by the BHF, that is in line with the overall objectives of the research group led by Doctor Matthew Stroud at the BHF Centre of Research Excellence, King’s College London (http://bit.ly/stroudlab). The aim of the project is to study the role of the nuclear envelope in cardiac development and disease using a range of molecular, cellular and physiological approaches. We are looking for a highly motivated researcher who possesses strong interpersonal skills with a biomedical background and will pursue collaborative research with members of the team. Furthermore, they will have expertise in cardiac development, physiology, cell biology, molecular biology. He/ she should also be a self-starter and able to work independently, accurately judge priorities, and have excellent organisational and communication skills. Previous experience of relevant techniques such as primary cell isolation and mass spectrometry will be considered highly suitable for this post. Knowledge and experience of integrative cardiac physiology approaches in animal models, such as in vivo cardiac phenotyping and management of gene-modified colonies will also be an advantage.
The BHF Centre of Research Excellence provides an excellent highly multidisciplinary environment with state-of-the-art equipment and facilities. Informal enquiries may be made to: matthew.stroud@kcl.ac.uk.
Developing brains use a mechanism like the Otoshi-buta (the drop lid), a kitchen wisdom.
Differentiating cells in embryonic cerebral walls form a dense filter-like layer to mechanically barrier nuclei of neural stem cells.
Loss of this barrier or fence results in abnormal popping out of neural stem cells’ nuclei, leading to inability of neural stem cells to produce new cells.
Brain formation relies on production of new cells by neural stem cells, which abundantly exist in the embryonic period. Neural stem cells are thin and elongated like radish sprouts (Kaiware-daikon) or Enoki mushrooms. They move their nuclei depending on the status of cell production. They divide along the inner (apical) surface of the wall and their nuclei are therefore near the surface just before and after mitosis, while their nuclei are away from the surface when they synthesize DNA in preparation for subsequent division. The range of this elevator-like (to-and-fro) nuclear movement is about 100μm even though the total length/height of each neural stem cell is ~300μm. Why (for which biological significance) it should be limited and how this range limitation is established are both unknown.
Research Results
In the present study, Professor Takaki Miyata, Assistant Professor Takumi Kawaue, and a 6th year medical student Yuto Watanabe in Nagoya University Graduate School of Medicine (dean: Kenji Kadomatsu, M.D., Ph. D.) showed that a transient layer consisting of differentiating cells and their dense processes mechanically barriers nuclei of neural stem cells. Experimental drilling of this barrier-like layer resulted in abnormal popping out of neural stem cells’ nuclei and the arrest of cell production by such nuclei-overshot stem cells. Thus demonstrated importance of mechanical limitation of nuclear movement during brain development is reminiscent of the usefulness of the Otoshibuta (Drop Lid) to limit a cooking space for condensing soap and avoiding undesired floating of ingredients.
Slides demonstrating the cooking analogy – click to expand
Research Summary and Future Perspective
Boomerangs or artificial satellites should turn back at an appropriate point. This study found that mouse neural stem cells’ nuclei start to come back after a 100-μm going because they are mechanically fenced by differentiating cells there. Drilling of the fence resulted in abnormal popping out of stem cells’ nuclei to 200μm. Since the normal range of nuclear shuttling is much greater in human (200μm), this study provides a basis of future comparative studies aiming at
elucidation of mechanisms underlying evolution of the human brain.
A postdoctoral position is available in the laboratory of Dr. Sophie Astrof at Thomas Jefferson University to study roles of cell-extracellular matrix (ECM) interactions in cardiovascular development and congenital heart disease. We have recently discovered that progenitors within the second heart field (SHF) give rise to endothelial cells composing pharyngeal arch arteries. Projects in the lab focus on the role of ECM in regulating the development of SHF-derived progenitors into endothelial cells and their morphogenesis into blood vessels. The successful candidate will combine genetic manipulation, embryology, cell biology, and confocal imaging to study molecular mechanisms of micro-environmental sensing during vascular development.
Astrof laboratory is a part of a modern and well-equipped Center for Translational Medicine at Jefferson Medical College (http://www.jefferson.edu/university/research/researcher/researcher-faculty/astrof-laboratory.html) located in the heart of Philadelphia.
To apply, please send a letter of interest detailing your expertise, CV and names and contact information of three references to sophie.astrof@gmail.com
A postdoctoral position is available in the lab of Thorold Theunissen at Washington University School of Medicine in St. Louis, Missouri, USA (theunissenlab.wustl.edu). Our research program is dedicated to exploring the molecular regulation of pluripotent stem cells and their applications in regenerative medicine. We have developed methods for inducing and maintaining human embryonic stem cells in a naive pluripotent state that shares defining transcriptional and epigenetic properties with the preimplantation embryo (Theunissen et al., Cell Stem Cell, 2014; Theunissen et al., Cell Stem Cell, 2016; Theunissen and Jaenisch, Development, 2017). We are seeking a highly motivated postdoctoral fellow who wishes to join a multidisciplinary research team investigating fundamental problems in stem cell biology. Projects are available in the following areas: (1) dissecting the signaling pathways and epigenetic mechanisms governing distinct human stem cell states, and (2) establishing novel stem cell models of early human development and disease. Assays used include RNA-seq, ChIP-seq, mass spectrometry, genome editing and flow cytometry.
Applicants should have a PhD and demonstrated relevant research experience. Excellent communication skills and the ability to work in collaboration are essential. A strong background in molecular biology, human ESC/iPSC culture, and differentiation assays is preferred. Experience in signal transduction, biochemistry and bioinformatics would be very beneficial.
Consistently ranked among the top 10 US medical schools, Washington University School of Medicine offers a highly interactive and stimulating academic environment for scientists in training. The candidate’s research will benefit from the highly collaborative environment within the Department of Developmental Biology and Center of Regenerative Medicine. We are located in the heart of the Central West End, a vibrant St. Louis neighborhood adjacent to major cultural institutions and one of the country’s largest urban parks. We offer competitive salary and benefit packages and candidates are eligible to apply for a Rita Levi-Montalcini Postdoctoral Fellowship offered by the Center of Regenerative Medicine.
To apply for this position please submit a CV, a cover letter describing research interests, and contact information for two references who can comment on your research to t.theunissen@wustl.edu. Applications will be reviewed promptly until the position is filled. Washington University is an equal opportunity employer and complies with applicable EEO and affirmative action regulations.
Hear from rodent and non-mammalian model organism users.
Learn about the highlight notice and applying for an NC3Rs grant.
Give a flash presentation: three minutes to speak about your research area, the specific expertise you offer and expertise you are seeking in a collaborator.
The topic: Segmentation is one of the most common features in animals and yet one of the most mysterious. The team led by Guillaume Balavoine explores the cellular and genetic bases of segment formation in the marine annelid Platynereis.
Segment formation in the annelid involves a special category of multipotent stem cells: teloblasts or posterior stem cells.
Platynereis is a very convenient model for microscopy and live-imaging. It is amenable to a number of genetic interference techniques such as morpholinos, RNAi and CRISPR-Cas9 knock-outs. Stable transgenesis can be performed using DNA transposition. We want to explore the biology of the teloblasts by developing a number of transgenic tools in Platynereis. We are performing clonal analysis using transgene mosaic insertions. We have developed a FUCCI tool to live-image the cell cycle transitions in individual cells.
Armed with these tools, we want to answer a number of questions concerning the stem cells: observe and characterize their lineage restrictions, observe asymmetric divisions and decipher the link between cell cycle and segment formation. Promising techniques such as light sheet microscopy, cell sorting and RNASeq analysis can be performed at the Institute.
The location: The Institut Jacques-Monod, funded jointly by the CNRS and the University Paris Diderot, is one of the main centers for basic research in biology in the Paris area. It is located near the center of Paris. The Institute provides an outstanding environment for research at the highest level and assembles around 30 groups working on genome and chromosome dynamics, cellular dynamics and signaling, development and evolution. The Institut Jacques Monod has developed a number of important core facilities, which all offer state-of-the art instrumentation and a high level of expertise. These include a large microscopy imaging platform and a genomics-transcriptomics facility.
Requirements: Candidate must have obtained PhD by the start date. Candidate will usefully have experience in developmental biology, including micro-injection, confocal imaging and image analysis. The position will remain open until filled.
Contact: To apply, please send your CV with a cover letter stating your motivation and contact details of three persons for references, to :
We are looking for a highly skilled and motivated candidate to join our group for a PostDoc position. In the Payer lab (http://www.crg.eu/bernhard_payer), we study epigenetic reprogramming in the mammalian germ line and the effects of ageing on fertility. In this project the prospective candidate will study the molecular links between ageing and oocyte quality decline in women. The work will be performed in collaboration with the fertility clinic Eugin in Barcelona (https://tinyurl.com/y9l834y5).
We are seeking applicants with a strong background in the fields of Mammalian Cell Culture, Embryology, Stem Cell Reprogramming and Differentiation, Tissue Engineering, Epigenetics and Reproduction. Excellent candidates from other related fields will also be considered.
Work Environment:
Our lab is part of the Gene Regulation, Stem Cells & Cancer Programme at the Centre for Genomic Regulation (CRG) in Barcelona, Spain (www.crg.eu). The CRG is a vibrant International Research Institute with Research Groups working in diverse fields such as Genomics, Cell and Developmental Biology, Systems Biology, Stem Cells, Cancer and Epigenetics. English is the working language.
Fellowship + Application Guidelines:
36 months by the INTREPiD Fellowship programme. Application guidelines including eligibility criteria can be found at: http://www.crg.eu/intrepid_fellowships
The application deadline is the 12th of October 2018, at 5:00pm (local time).
For informal questions regarding the position, please contact: academicoffice@crg.eu
It’s undoubtedly the middle of summer here in Saint Augustine, Florida. Daily temperatures are soaring into the 90s, and we’re grateful if the humidity dips below 70%. Thankfully, the Seaver lab doesn’t have to contend with much of this heat. Instead, our members are inside, comfortable though busier than ever, mentoring summer interns, piloting new experiments, and writing up papers. It’s summertime, and the University of Florida’s Whitney Laboratory for Marine Bioscience is bustling with research as a whole!
I am Alexis Lanza, a PhD candidate in Dr. Elaine Seaver’s Lab with a research interest in embryonic signaling events critical for dorsal ventral axis formation in annelids. Here, I share a bit about our lab and its daily activities.
Dr. Seaver and graduate student Alexis Lanza imaging samples using a compound microscope
Area of Research
Broadly speaking, the research being done in the Seaver lab aims at answering questions about the development and evolution of marine invertebrates. We’re curious about the cellular and molecular mechanisms that drive developmental processes from an unfertilized egg through to zygotic, larval, and juvenile stages, as well as during regeneration. We’re also interested in how molecular differences that exist across species may account for body plan differences across animals. In our lab we use a small marine worm called Capitella teleta as our model organism.
Capitella teleta as an ideal model
Adult Capitella teleta.
Capitella teleta is a polychaete annelid worm and member of the
Lophotrochozoa. It is a cosmopolitan species and can be found along intertidal zones living in organically rich marine sediments. Capitella can easily be kept as part of a lab colony in glass finger bowls with seawater and roughly a tablespoon of mud. Following fertilization, its embryos develop into pelagic non-feeding larvae that metamorphose into juvenile worms approximately 9 days post-fertilization. At around 8 weeks old, these juveniles mature into adults that are easily distinguishable by sex and are capable of reproducing year-round. Capitella’s ability to produce embryos regularly is one reason why it is an ideal candidate for early developmental studies.
Furthermore, its embryos undergo a stereotypic cleavage program called unequal spiral cleavage in which blastomere formation occurs according to a predictable order, size, and position. This predictable cleavage program is also shared by other taxa including mollusks and nemerteans, and allows for the identification of individual blastomeres that can then be microinjected, or deleted using a laser (Meyer and Seaver, 2010; Yamaguchi et al., 2016).
Early cleavage stage embryos of Capitella teleta
Another prominent area of study within our lab focuses on how regeneration occurs in Capitella (de Jong and Seaver, 2017). Capitella, like many of its annelid counterparts, possesses regenerative capabilities following transverse amputations, as it is capable of regenerating posterior segments. However, Capitella cannot regenerate anteriorly. The development of Capitella as an annelid model for developmental and regeneration studies was rooted most significantly in our ability to conduct experimental manipulations, functional genomics, as well as having a sequenced, annotated, and slowly evolving genome (Simakov et al., 2012). Altogether, the Seaver lab utilizes tools of experimental embryology, molecular and cellular biology, and functional genomics to answer evo/devo questions.
A typical day of animal care
Maintaining a Capitella colony requires several activities including feeding. While these tasks aren’t difficult in themselves, every member of the Seaver lab chips in throughout the week, tackling a different task during breaks in experiments.
We keep our Capitella lab colony in glass finger bowls in a 19°C incubator
Sifting
Every glass finger bowl initially contains 40 individual worms. As the worms become reproductive however, we need to make an effort to avoid overpopulation in each bowl. When “sifting,” worms are removed from their bowl of mud and carefully sorted under a dissecting microscope. Unhealthy worms and larvae (the product of spontaneous matings) are removed, and only healthy individuals are placed into a fresh bowl of mud to be restocked as part of the working colony used for experiments.
Worms are sorted under a dissecting microscope and healthy individuals are placed into a fresh bowl of mud.
The Seaver lab divides its colony of bowls into two groups, each of which is sifted on an alternating, bi-weekly schedule. At the start of each week, a table listing all the colony bowls that need to be sifted that week is made.
Every week, lab members sign up to sift a couple colony bowls.
Lab members will then sign up for particular bowls based on their experimental needs are for the week. For instance, colony bowls with worms that have only recently reached sexual maturity are less likely to have larvae than would older bowls. Thus, if aiming to collect larvae one might want to sift an “older bowl.” Alternatively, if the goal is to conduct embryological experiments one may want to collect mature male and female worms, which can be used as a mating pair to produce early stage embryos.
Setting up the next generation
To maintain our lab colony through generations, it’s crucial that we plan ahead. As our worms age, they become less healthy and produce less offspring so are gradually phased out via the sifting process. To account for losing these individuals we introduce young, sexually mature worms (approx. 8 weeks old), all of which were raised from larval stages.
During the sifting process, lab members collect late stage larvae (approx. 8-9 days old), which are then used to create the new generation of colony bowls. First, glass fingerbowls are filled with seawater plus a teaspoon sized scoop of mud. Next, late stage larvae from several different mating pairs are pipetted into said fingerbowls until the total number of larvae equals 40. The mud is used to induce larvae to metamorphose. These new bowls will then be raised at 16°C for 8 weeks before being incorporated into the working colony.
Feeding
Animals need to be fed fresh mud once a week. We collect, sieve and freeze large quantities of marine mud that can then be thawed and used for animal care over the course of several months. To feed the colony, seawater is first decanted from each bowl. One tablespoon of mud is then added to each bowl, followed by the addition of fresh seawater. This process takes anywhere from 10 minutes to 1 hour depending on how large the colony is at any given time.
Fingers crossed for embryos
Mating Bowls
Remember those mature male and female worms collected during the sifting process? To acquire early cleavage stage embryos, mature male and female worms are separated by sex into bowls with 4 or 5 individuals each for about 3 days. When ready to set up a mating we simply combine the individuals from a male bowl with those from a female bowl. In general, we’ve noticed that it takes about 8-10 hours for the worms to mate and the females to lay their broods. Since most of our embryology experiments begin in the morning we need to set up these matings around midnight… this often means taking your worms home with you and only giving vague answers to questions your housemates may ask.
In the morning, back at the lab, we sift our mating bowls with bated breath to check if any of the female worms laid a brood of fertilized embryos.
When female worms are ready to lay their eggs, they create an encasement called a brood tube. The worm remains inside the brood tube along with her offspring until they become larval stages.
Using our dissecting microscopes and a pair of forceps, we dissect open any brood tubes and gently pipette the embryos into a clean petri dish. From here we can stage the embryos and determine how many cleavage divisions into development they are. Capitella embryos are approximately 200 microns in size and each cleavage division takes approximately 45 minutes. These embryos are amenable to chemical perturbants, allowing us to manipulate various signaling pathways and determine its role in development (Lanza and Seaver, 2018). Furthermore, Capitella embryos can be microinjected. To do so, the egg membrane of the embryo is first softened using a sucrose: sodium citrate mixture. This treatment is typically performed one cleavage division before the desired cell stage you wish to inject. We can then microinject individual cells during early cleavage stages with aqueous or lipophilic lineage tracers, DNA, RNA, or morpholinos, and then raise these embryos in seawater to larval stages (Meyer et al., 2010; Seaver, 2016). Like in many other species, most of the difficulty about working with early cleavage stage embryos is timing related. Since we can’t predict exactly when the eggs are fertilized we often end up racing against the clock to get our experiments underway before our target cleavage cycle ends or wait for hours until our embryos finally make it to the target cell stage. In general, after experimental manipulation, embryos are raised for six days to larval stage before being fixed for phenotypic analysis.
By the day’s end, the afternoon showers have usually subsided and the temperature has waned. It’s around then that the members of the Seaver lab emerge – just in time to enjoy the last few hours of sunlight during another Florida summer.
Members of the Seaver lab Spring 2018. Left to right, Linlin Zhang, Elaine Seaver, Alexis Lanza, Stephanie Neal.
The School of Natural Sciences invites applications for three full-time, permanent Lectureships in the broad area of Biology encompassing the full spectrum of genes, organisms and ecosystems. As part of the School of Natural Sciences, the biology and zoology subject areas are dynamic and growing elements with strong research records and successful recruitment onto our undergraduate and postgraduate degrees. Currently we are looking to expand our research base and our teaching and outreach capabilities. Bangor University is committed to excellence in research, teaching & scholarship and offers attractive career prospects for staff with such expertise.
Details of current staff research interests are available online. The School benefits from modern research laboratories and microscopy suites, a Natural History Museum, freshwater and marine aquaria, fly room, reptile and rodent facilities, plus a number of temperature controlled rooms, a botanical garden located on the fringes of the Menai Strait, a herbarium, and avian housing facility. Further research opportunities are facilitated by close links to the Centre for Ecology and Hydrology (CEH), the Centre for Environmental Biotechnology (CEB), the Schools of Ocean Sciences, Computer Sciences, and Psychology and access to the Supercomputing Wales High Performance Computing cluster.
Candidates should have a PhD in biological science, be able to develop research programmes of their own, contribute to the School’s Research Excellence Framework (REF) submission, show an enthusiasm for teaching, be willing to engage with outreach activities and also have excellent interpersonal skills.
Interviews will be held in the week commencing 3rd September. The successful candidates will be expected to start on 1st October 2018 or as soon as possible thereafter.
Informal enquiries may be made to Professor Chris Freeman, email c.freeman@bangor.ac.uk; and copied to Dr Nia Whiteley, email n.m.whiteley@bangor.ac.uk.
The transience of flowers is proverbial. Degeneration of flowers is elicited after successful pollination by the onset of seed and fruit development. However, also unpollinated flowers do not last forever – on the contrary, the life span of unpollinated flowers is a tightly regulated trait that differs greatly among plant species. Some plant species like orchids of the genus Phalenopsis have very long-lived flowers that can stay receptive for weeks, even months. Most plant species, however, sustain unpollinated flowers for much shorter time spans. In the most extreme cases mere hours pass between flower opening and withering.
In the Programmed Cell Death lab at the VIB Center for Plant Systems Biologywe are using the model plant Arabidopsis thaliana to study the longevity and senescence of unpollinated flowers. Although Arabidopsis is the most advanced model species for plant developmental biology, it is actually not particularly convenient to investigate the biology of flower senescence. Arabidopsis flowers are tiny, and moreover they are self-pollinated, so that the male floral parts (anthers) have to be either removed manually by emasculation, or male-sterile mutants have to be used.
A typical Arabidopsis flower aligned next to a pin. The floral stigma is at the level of the pointed end of the needle.
Every Arabidopsis researcher knows how to perform emasculation and pollination in order to cross different mutants, marker lines, or accessions. And many of us know that a flower emasculated on Friday might already have started to senesce by Monday, hence making successful pollination and seed set inefficient or impossible.
In order to observe and stage flower withering in Arabidopsis, we set up a macroscopic imaging system using a single lens reflex camera equipped with a macro objective. Taking images every 10 minutes under constant light enabled us to monitor flower senescence in detail. We could observe that concomitant with the withering of petals and sepals, also the floral stigma was degenerating.
The stigma of flowers is the primary receptive surface for pollen grains. In Arabidopsis the stigma consists of over 200 elongated fingerlike cells, the so-called stigmatic papillae, specialized epidermal cells that serve to intercept the pollen. In Arabidopsis, hundreds of pollen grains compete to fertilize about 50 ovules located inside the pistil of each flower. Once in contact with the stigma cells, pollen grains will hydrate, and germinate to form a single tip growing cell, the pollen tube. The pollen tube penetrates the papilla cell wall, growing in between the cell wall and the plasma membrane to the base of the papilla cell. From there on, guided by chemical cues, the pollen tube grows through the style and the transmitting tract to the ovules. There, it will release its two sperm cells to fertilize the egg cell and the central cell of the female gametophyte, thus initiating embryo and seed development.
The journey of the pollen tube during plant reproduction in Arabidopsis. Pollen grains (orange) are shed from the anthers and adhere on the stigmatic papilla cells. During the preovular guidance phase, pollen grains hydrate and germinate, and pollen tubes (red) penetrate in the papilla cells and grow through the style and transmitting tract. After that, pollen tubes are attracted by the ovules, enter the ovules with a near one-to-one relationship during the ovular guidance phase. Finally, pollen tube growth stops, and the sperm cells are released into the female gametophyte to perform double fertilization. Picture is a courtesy of Zhen Gao.
The structure of the intact Arabidopsis stigma appears a bit like a little hedgehog due to the hundreds of erect elongated papilla cells. Once flower senescence sets in, individual papilla cells start to break down, leading first to a ragged, and finally to a completely collapsed appearance of the stigma.
Pollination assays at different time points during this process showed that stigma degeneration coincided with a sharp decline in seed set, suggesting that viable papilla cells are necessary for successful reproduction. To test this hypothesis, we specifically ablated papilla cells by expressing diphtheria toxin chain A (DT-A) under a stigma-specific promoter. As we had assumed, DT-A induced stigma degeneration likewise caused a strong reduction of seed set.
In order to quantify stigma longevity, we faced a considerable problem – the macro setup was only able to follow a single flower at a time, and a price tag of about 1400 Euro per unit made the parallel acquisition of time courses impracticable. Coincidence came to our aid: Setting up a skype conference call we noticed that a cheap webcam without autofocus can be brought very close to an object (in that particular case a certain PhD student’s uvula) and amazing magnification can be achieved. After some experimentation with different webcams, we rigged up a phenotyping system with 20 webcams. We used a custom-made script to operate the open source camera surveillance software so that each camera acquires one picture frame every 10 minutes. Although the image quality was of course not comparable to the conventional macro lens setup, in the end of the day we had established a phenotyping platform with 20 imaging units for mere 250 Euros.
Webcam imaging platform to follow flower senescence in Arabidopsis
Analysis of the time lapse images revealed a remarkable reproducibility of stigma collapse on average around 56 hours after emasculation of a flower at the developmental stage 12c1.
The apparently precisely timed collapse of papilla cells suggested that an active cell death process might be occurring in Arabidopsis stigmata. Based on a large-scale meta-analysis of mRNA transcriptome profiles, our lab has established promoter-reporter lines of genes transcriptionally upregulated prior to a number of developmentally regulated programmed cell death (PCD) events2. Microscopic analyses revealed that most promoter-reporters come up during stigma senescence, thus linking stigma degeneration with established forms of developmentally controlled PCD processes3. Microscopic imaging of papilla cells is not trivial; although the individual cells are thankful objects for cell biological analyses, the stigma as a whole is a rather large and very sensitive three-dimensional structure. In order to investigate cell biological hallmarks of PCD in living stigmata, we could not use conventional slide-and-coverslip setups lest we create injuries and mounting artefacts in papilla cells. After testing diverse setups, we found that mounting the entire flower in an agar block and viewing the stigma from top with a long working-distance water dipping lens on an upright Zeiss 710 confocal microscope allowed a minimally invasive way to perform live-cell imaging of papilla cell death. In order to visualize cell death, we used a live/death stain in which fluorescein-diacetate (FDA) fluorescence indicates living cells, while nuclear staining of the membrane impermeable stain propidium iodide (PI) reveals plasma-membrane permeation as a committing step of cell death.
Cell death/ viability staining of stigmatic papilla cells. Fluorescein-diacetate (FDA) fluorescence indicates living cells, while nuclear staining of the membrane impermeable stain propidium iodide (PI) reveals plasma-membrane permeation as a committing step of cell death. Picture is a courtesy of Yulia Salanenka.
Alternatively, we used a tool developed in our lab, a tonoplast integrity marker (ToIM) line4. This plant line expresses a cytoplasmic green fluorescent protein (cGFP) and a polycistronically produced vacuolar red fluorescent protein (vRFP). Living cells display clearly separated GFP and RFP domains indicating an intact vacuolar membrane (tonoplast). Collapse of the large central vacuole in mature plant cells, another hallmark of plant programmed cell death5, is visible as merging of red and green fluorescence signals.
A confocal maximum intensity projection of a representative stigma from a Tonoplast Integrity Marker line (ToIM). Green labels cytosolic GFP, red shows RFP localized to the vacuole. The fusion of both signals appears yellow and indicates a loss of cellular compartmentalization symptomatic of cell death.
Our analyses revealed that as in the macroscopic setup, stigma degeneration occurred over a time span of 12 hours. Fascinatingly, the death of individual papilla cells followed a pattern from the periphery towards the center of the stigma, and often small clusters of individual cells underwent vacuolar collapse almost synchronously. On the level of individual cells, the PCD program occurred surprisingly fast; vacuolar collapse, plasma membrane permeation, nuclear fragmentation, and finally cellular collapse occurred within about one hour. The reproducibility of these successive events confirmed our hypothesis of a tightly regulated cell death program.
In order to discover regulators of this cell death process, we set out to perform an RNA-seq analysis monitoring the transcriptional changes occurring in the stigma over time. We manually dissected the stigmata from flowers at 1, 2, and 3 days after emasculation, corresponding to young, mature, and senescent flower stages. In total we had to stage, emasculate, and dissect way over individual 2000 flowers, a feat that we only managed with lots of training and a great team effort.
Emasculation session in the PCD lab. The combined efforts of at least three people were required during the laborious process of Arabidopsis flower emasculation. The stigmata of about 2000 emasculated flowers was collected and subjected to RNA-Seq analysis.
Illumina RNA-seq revealed a large number of differentiallty regulated genes (1180 genes out of more than 25 000 predicted genes in Arabidopsis) over the course of stigma senescence. Interestingly far more genes were upregulated (897) than downregulated (283), again indicating an actively controlled program controlling stigma senescence and papilla cell death.
RNA sequencing also confirmed the strong upregulation of developmental PCD-associated genes. In order to identify key regulators of this process, we investigated the expression profiles of transcription factors (TFs). Interestingly, the plant specific NAC TF family was strongly overrepresented in senescence-associated TFs. Among the most strongly upregulated NAC TFs was the well-established leaf senescence regulator ORESARA1/ANAC092 (ORE1, Korean for “long-living”), and the flower-specific ANAC074, a previously uncharacterized NAC TF that we dubbed KIRA1 (KIR1) after the killer “Kira” in the Japanese manga “Death Note”.
Using our webcam-phenotyping platform, we investigated an established knock-out mutant of ORE1 and found a slight, but not significant delay of stigma collapse. Interestingly, a newly established KIR1 knock-out mutant showed modest, but significant extension of stigma life span. We crossed the mutants and the resulting kir1 ore1 double mutant showed a clear synergistic effect which led to a doubled life span in comparison to the wild type. We also generated lines overexpressing dominant repressive mutant versions of KIR1 and ORE1, and some of these lines had an even stronger effect with some stigmata only collapsing at 11 days after emasculation. Intriguingly, both recessive kir1 ore1 loss-of-function mutations, as well as expression of dominant repressive alleles showed an uncoupling of stigma life span from floral organ longevity. Although the stigma life span was considerably increased, sepals and petals senesced similar to the wild type.
In a complementary approach, we investigated KIR1 gain-of-function mutants. Estradiol-inducible systemic misexpression of KIR1 in seedlings caused a rapid growth arrest caused by a widespread ectopic cell death. This demonstrates that the transcriptional program controlled by KIR1 is sufficient to elicit cell death outside of the stigma context. When inducing precipitate KIR1 expression specifically in the stigma, the papilla cell death and loss of receptivity occurred significantly earlier than in estradiol treated wild types or mock treated mutant lines. These results demonstrated that KIR1 functions to actively terminate the receptive life span of the flower by promoting a cell death program in the stigma.
This picture shows an emasculated Arabidopsis flower undergoing localized estradiol treatment in order to induce KIR1-GFP overexpression in the stigma. The floral stigma is imbibed into an estradiol-containing droplet for 6 hours and pollinated after that.
A central question arising from this investigation was of course whether aged but viable kir1 ore1 loss-of-function mutants could be successfully pollinated. When performing a pollination time series, we discovered that in dominant repressive KIR1 and ORE1 mutants, there was as significant, but rather modest extension of flower receptivity. In the recessive kir1 ore1 double mutant, there was no extension of floral receptivity at all. We used pollen from a transgenic Arabidopsis line that expresses β-glucuronidase (GUS) under a pollen-tube specific promoter to visualize pollen tube growth on pollinated stigmata after addition of the GUS substrate X-Gluc6. Microscopic analyses showed that pollen hydrated and germinated, but that pollen tube growth on aged mutant stigmata was strongly reduced in comparison to younger wild type or mutant stigmata. No pollen tubes could be detected to enter the style or the transmitting tract of the aged mutant flowers, suggesting that KIR1 / ORE1 loss of function is sufficient to suppress age-induced stigma PCD, but not sufficient to maintain stigma function in a corresponding fashion.
In summary, our research on stigma senescence suggests that a KIR1-ORE1 dependent cell death program actively terminates stigma and flower receptivity. However, as suppression of this cell death program is not sufficient to extend stigma function, we assume there must be additional, KIR1 – ORE1 independent pathways that either passively or actively terminate stigma function in the absence of cell death. With research going on in our lab we attempt to research these pathways in order to effectively modulate flower receptivity.
While Arabidopsis serves as a model system to discover these pathways, our research might also be applicable to outcrossing crop species. An extension of floral receptivity, especially under spells of environmental stresses, which are deleterious to plant reproduction, might be a key strategy to stabilize the yield of seed and fruit bearing crops7.
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