One of the central models for understanding the long-range coordination of pattern formation and growth during animal development is the Decapentaplegic/Bone Morphogenetic Protein (DPP/BMP) morphogen gradient in the developing Drosophila wing.  During development, the DPP gradient is thought to serve two primary functions: controlling the concentration dependent patterning of target gene expression as well as directing uniform cell proliferation. Upon initiating the lab in 2006, we used an unbiased microarray approach to identify novel gene targets of this signaling pathway, with an eye toward the identification of factors encoding downstream targets involved in either growth or epithelial morphogenesis.  The first target that we characterized encodes the novel extracellular protein Larval Translucida (LTL), which we found to be a feedback modulator of DPP/BMP activity (Szuperak et al., 2011). To build on these studies, we next undertook independent efforts to 1) directly visualize DPP in vivo using novel antibody reagents that have long eluded the field and 2) exploit the emerging CRISPR/Cas9 technology to test the precise requirements for the DPP morphogen gradient in both pattern formation and growth control.  Toward the first aim, we successfully generated a polyclonal antibody that detects the non-secreted pro-domain of the DPP protein in vivo.  We analyzed hypomorphic mutant clones to validate the antibody‚Äôs specificity, and these experiments led to the unexpected finding that large hypomorphic clones lacking normal DPP expression did not strongly perturb disc growth.  To circumvent the difficulty of working with haploinsufficient DPP null alleles and directly test the requirements for the DPP gradient in disc growth, we employed CRISPR/Cas9 to engineer a novel conditionally-inducible DPP null allele that allows for precise control over gene expression in vivo.  Using this allele to inactivate DPP in its endogenous compartmental stripe domain, we demonstrated that a gradient of DPP signaling was essential for patterning but dispensable for uniform proliferation (Akiyama and Gibson, Nature 2015).


The organization of cells into polarized and adherent epithelia layers is a core feature of animal development, yet we still know relatively little about the cell biology of mitosis in the epithelial context. Indeed, while the fundamental cell biology of mitosis is well studied in yeast, tissue culture cells, and early embryos, precisely how cell division is executed within the spatial and mechanical constraints of polarized epithelia has received far less attention.  A key outcome of our previous work was the finding that pseudostratified epithelial cell division in Drosophila wing discs and Nematostella larvae is descriptively similar, if not identical, to the process of interkinetic nuclear migration (IKNM) as described in vertebrate neuroepithelia.  In brief, a universal or near-universal feature of pseudostratified epithelia is that the nuclei of proliferating cells oscillate between medial positions in interphase and apical positions during mitosis, when cells undergo extensive cytoskeletal rearrangements and round up at the apical surface of the epithelium.  A further conserved feature of apical rounding is the orientation of the mitotic spindle to the plane of the epithelium, a phenomenon we term planar orientation.  The mechanism by which the mitotic spindle machinery interacts with elements of the cell polarity apparatus to achieve planar orientation has remained poorly understood.  To address this problem, we recently established a detailed description and time course for cytoskeletal events, which led to the observation that the mitotic spindle poles align precisely to the plane defined by the position of the basolateral septate junctions.  By disrupting the basolateral tumor suppressor proteins Discs Large (DLG) and Scribbled (SCRIB), we observed that a primary phenotypic consequence was the misorientation of the mitotic spindle.  We next investigated the consequences of spindle misorientation for tissue architecture and found that the basal daughters of aberrant divisions frequently delaminated from the epithelium and subsequently underwent apoptotic cell death.  Intriguingly, when cell death was inhibited, the delaminated cells accumulated basally and exhibited features of epithelial-to-mesenchymal transition (EMT), including loss of E-Cadherin and upregulation of matrix metalloproteases.  Taken together, this work has uncovered the molecular basis for planar spindle orientation in proliferating epithelia and has allowed us to propose a new model for how defects in epithelial cell division could lead to EMT events during human disease (Nakajima et al., Nature 2013). 

In a parallel set of experiments designed to discover new factors implicated in mitotic control, we used a FACS/microarray approach to globally define cell cycle-associated transcriptional oscillations in vivo.  We then compared the transcriptional oscillations observed in wing disc cells with those found in Drosophila S2 cells and found a surprising degree of context-specific transcriptional regulation, even for some of the basal components of the DNA replication machinery.  Extending these results, we performed RNAi knockdown on several hundred cyclically expressed genes, which identified over 100 cyclic genes required for wing growth.  We further characterized the function of the periodic genes required for wing growth with a secondary FACS screen for defects in cell cycle phasing as well as a confocal screen for visible mitotic defects and visible defects in interkinetic nuclear migration. Strikingly, many of the cyclic genes required for cell cycle progression or mitosis were not identified in prior cell-based RNAi screens.  In sum, this work not only provided a genome-wide perspective on cyclic transcription, but also identified and functionally validated numerous novel regulators of cell proliferation in vivo (Liang et al., Developmental Cell 2014).


A third area of increasing emphasis in our lab has been the use of the starlet sea anemone Nematostella vectensis as a genetically tractable model for understanding epithelial organization and developmental cell biology in a representative early-branching metazoan.  Nematostella occupies an ideal phylogenetic position for understanding the evolutionary history of traits within Bilateria.  The animals are easy to culture in the lab, produce vast numbers of embryos after an induced spawning, and feature rapid development to the juvenile stage.  We have established a colony of approximately 8,000 individuals, 3,000 of which have been sex-segregated into spawning groups, which allows us to systematically obtain hundreds of embryos twice per day, five days per week.  In the last four years we have initiated several new projects still in progress, and have completed a detailed analysis of tentacle patterning and morphogenesis in Nematostella (Fritz et al., Development 2013).  Through these efforts we have established methods for confocal and light microscopy, embryo injection, and custom microarray and mRNA-seq analyses.  These studies set the stage for future work on a broad range of questions about Nematostella development and motivated us to generate genetic tools akin to those we employ in Drosophila.  In this regard, the most significant accomplishment in our Nematostella work in the last five years was the validation of TALEN and CRISPR/Cas9-mediated homologous recombination.  We first validated TALEN and CRISPR/Cas9-based mutagenesis protocols by knocking out the endogenous Nematostella fluorescent protein encoded by NvFP-7R.  We next established conditions for homologous recombination of DNA constructs into the NvFP-7R locus, thus creating a transgenesis method that exploits the visible loss of NvFP-7R expression as a highly efficient readout for transgene insertion.  Although almost purely technical in nature, this work will open the door for a new generation of molecular-genetic manipulations never before possible in pre-bilaterian phyla (Ikmi et al., Nature Communications 2015).


Gibson Lab

1000 East 50th Street

Kansas City, MO 64110