Role of site-directed mutagenesis and adjuvants in the stability and potency of anthrax protective antigen

Anthrax is a zoonotic infection caused by the gram-positive, aerobic, spore-forming bacterium Bacillus anthracis. Depending on the origin of the infection, serious health problems or mortality is possible. The virulence of B. anthracis is reliant on three pathogenic factors, which are secreted upon infection: protective antigen (PA), lethal factor (LF), and edema factor (EF). Systemic illness results from LF and EF entering cells through the formation of a complex with the heptameric form of PA, bound to the membrane of infected cells through its receptor. The currently available anthrax vaccines have multiple drawbacks, and recombinant PA is considered a promising second-generation vaccine candidate. However, the inherent chemical instability of PA through Asn deamidation at multiple sites prevents its use after long-term storage owing to loss of potency. Moreover, there is a distinct possibility of B. anthracis being used as a bioweapon; thus, the developed vaccine should remain efficacious and stable over the long-term. Second-generation anthrax vaccines with appropriate adjuvant formulations for enhanced immunogenicity and safety are desired. In this article, using protein engineering approaches, we have reviewed the stabilization of anthrax vaccine candidates that are currently licensed or under preclinical and clinical trials. We have also proposed a formulation to enhance recombinant PA vaccine potency via adjuvant formulation.     

Anomaly of cornea and ocular adnexa in spinster homolog 2 (Spns2) knockout mice

Spinster 2 (Spns2) is a transporter that pumps sphingosine-1-phosphate (S1P), a bioactive lipid mediator synthesized in the cytoplasm, out of cells into the inter cellular space. S1P is a signal that modulates cellular behavior during embryonic development, inflammation and tissue repair, etc. A Spns2-null (KO) mouse is born with failure of eyelid closure (eyelid-open-at birth; EOB) and develop corneal fibrosis in adulthood. It remains elusive whether corneal lesion is caused by exposure to keratitis (lagophthalmos) of EOB phenotype or the loss of Spns2 directly perturbs the corneal tissue morphogenesis and intra-eyelid structures. Therefore, we investigated differences between the cornea and ocular adnexa morphogenesis in KO and wild-type (WT) embryos and adults as well.The loss of Spns2 perturbs cornea morphogenesis during embryonic development as early as E16.5 besides EOB phenotype. Histology showed that the corneal stroma was thinner with less extracellular matrix accumulation, e.g., collagen and keratocan in the KO mouse. Epithelial stratification, expression of keratin 12 and formation of desmosomes and hemidesmosomes were also perturbed in these KO corneas. Lacking Spns2 impaired morphogenesis of the Meibomian glands and of orbicularis oculi muscles. KO glands were labeled for ELOVL4 and PPARγ and were Oil-Red O-positive, suggesting KO acinar cells possessed functionality as the glands.This is the first report on the roles of Spns2 in corneal and Meibomian gland morphogenesis. Corneal tissue destruction in an adult KO mouse might be due to not only lagophthalmos but also to an impaired morphogenesis of cornea, Meibomian glands, and orbicularis oculi muscle.     

Hedgehog signaling regulates dental papilla formation and tooth size during zebrafish odontogenesis

Background : Intercellular communication by the hedgehog cell signaling pathway is necessary for tooth development throughout the vertebrates, but it remains unclear which specific developmental signals control cell behavior at different stages of odontogenesis. To address this issue, we have manipulated hedgehog activity during zebrafish tooth development and visualized the results using confocal microscopy. Results : We first established that reporter lines for dlx2b, fli1, NF‐κB, and prdm1a are markers for specific subsets of tooth germ tissues. We then blocked hedgehog signaling with cyclopamine and observed a reduction or elimination of the cranial neural crest derived dental papilla, which normally contains the cells that later give rise to dentin‐producing odontoblasts. Upon further investigation, we observed that the dental papilla begins to form and then regresses in the absence of hedgehog signaling, through a mechanism unrelated to cell proliferation or apoptosis. We also found evidence of an isometric reduction in tooth size that correlates with the time of earliest hedgehog inhibition. Conclusions : We hypothesize that these results reveal a previously uncharacterized function of hedgehog signaling during tooth morphogenesis, regulating the number of cells in the dental papilla and thereby controlling tooth size. Developmental Dynamics 244:577–590, 2015. © 2015 Wiley Periodicals, Inc.     

The life cycles of two species of Myxomycetes in Physarales,Physarum rigidum and Didymium squamulosum

Myxomycetes are eukaryotic microorganisms containing characteristics akin to both fungi and amoebae. They can complete their whole life cycles while being cultured on agar media, and under‐laboratory conditions, which favors taxonomic, phylogenetic, and cytological researches. Here, we describe the life cycles of two such species: Didymium squamulosum collected from the field and Physarum rigidum cultured from moist chamber both belonging to the Order Physarales. Three per cent oat‐agar media (OAM) was used to culture the plasmodia until they aggregated and were almost starved. Natural light was then applied to the plasmodia to induce fructification. Their life cycles share the same common stages, namely: spore, myxamoebae, swarm cell, plasmodia, and sporulation. In this study, we describe the morphogenesis from spore to spore of two species by differential interference contrast (DIC) and stereoscopic microscopies, as well as discuss the differences between the development of both species and interspecies. We found that the spore germination method of both species was the same. However, there were differences noted in time taken and fruiting body formation. Unlike P. rigidum, the species D. squamulosum did not require natural light stimulation. Moreover, the maturation process of both species had similar color transitions but exhibited distinct morphology in each developmental stage except during the swarm cell stage.     

Response of Escherichia coli growth rate to osmotic shock

It has long been proposed that turgor pressure plays an essential role during bacterial growth by driving mechanical expansion of the cell wall. This hypothesis is based on analogy to plant cells, for which this mechanism has been established, and on experiments in which the growth rate of bacterial cultures was observed to decrease as the osmolarity of the growth medium was increased. To distinguish the effect of turgor pressure from pressure-independent effects that osmolarity might have on cell growth, we monitored the elongation of single Escherichia coli cells while rapidly changing the osmolarity of their media. By plasmolyzing cells, we found that cell-wall elastic strain did not scale with growth rate, suggesting that pressure does not drive cell-wall expansion. Furthermore, in response to hyper- and hypoosmotic shock, E. coli cells resumed their preshock growth rate and relaxed to their steady-state rate after several minutes, demonstrating that osmolarity modulates growth rate slowly, independently of pressure. Oscillatory hyperosmotic shock revealed that although plasmolysis slowed cell elongation, the cells nevertheless “stored” growth such that once turgor was reestablished the cells elongated to the length that they would have attained had they never been plasmolyzed. Finally, MreB dynamics were unaffected by osmotic shock. These results reveal the simple nature of E. coli cell-wall expansion: that the rate of expansion is determined by the rate of peptidoglycan insertion and insertion is not directly dependent on turgor pressure, but that pressure does play a basic role whereby it enables full extension of recently inserted peptidoglycan.Cell growth is the result of a complex system of biochemical processes and mechanical forces. For bacterial, plant, and fungal cells, growth requires both the synthesis of cytoplasmic components and the expansion of the cell wall, a stiff polymeric network that encloses these cells. It is well established that plant cells use turgor pressure, the outward normal force exerted by the cytoplasm on the cell wall, to drive mechanical expansion of the cell wall during growth (1, 2). In contrast, our understanding of the physical mechanisms of cell-wall expansion in bacteria is limited. Furthermore, the bacterial cell wall is distinct from its eukaryotic counterparts in both ultrastructure and chemical composition. In particular, the peptidoglycan cell wall of Gram-negative bacteria is extremely thin, comprising perhaps a molecular monolayer (3). This raises the question of whether these organisms require turgor pressure for cell-wall expansion, or whether they use a different strategy than organisms with thicker walls.Turgor pressure is established within cells according to the Morse equation, P = RT(C_in ? C_out), where C_in is the osmolarity of the cytoplasm, C_out is the osmolarity of the extracellular medium, R is the gas constant, and T is the temperature. In the Gram-negative bacterium Escherichia coli, P has been estimated to be 1–3 atm (4, 5). A primary role of the cell wall is to bear this load by balancing it with mechanical stress, thereby preventing cell lysis. In 1924, Walter proposed a theory of bacterial growth based on the premise that mechanical stress, in turn, is responsible for stretching the cell wall during growth (6). In support of this theory, he and others showed that the growth rate of a number of bacterial species, including E. coli, decreases as the osmolarity of their growth medium is increased (6–9) (Fig. 1A). This result would be predicted by the Morse equation if growth rate, e˙, scaled with pressure: e˙∼P∼−Cout. The decrease in growth rate did not depend on the chemical used to modulate the osmolarity, which demonstrated that this effect was not due to specific, toxic reactions. More recently, several theoretical studies have offered plausible molecular mechanisms by which turgor pressure could drive cell-wall expansion. These theories range from ones in which the cell wall is irreversibly stretched by turgor pressure when peptidoglycan cross-links are hydrolyzed (10, 11) to ones in which the rate of peptidoglycan biosynthesis is directly dependent on turgor pressure (12, 13). Two of these theories explicitly predict the scaling between cellular growth rate and pressure (12, 13), providing possible explanations of the classic experiments (6–9).Open in a separate windowFig. 1.E. coli cells maintain their elongation rate after hyperosmotic shock. (A) The population-averaged steady-state elongation rate as a function of the concentration of sorbitol in the growth medium, C_out (red circles). Also shown are the population-averaged elongation rates during phase III of recovery from a 400 mM hyperosmotic shock (blue triangles) and immediately after a 400 mM hypoosmotic shock (green squares). Each data point is averaged over 20–100 cells. Error bars indicate ±1 SE. (Inset) Definition of elongation rate. (B) E. coli stained with fluorescent WGA, imaged with phase and epifluorescence microscopy. (Lower) Schematic defining the length and width of a rod-shaped cell. (C) Longitudinal strain, ε_l, and radial strain, ε_w, as a function of medium osmolarity. Error bars indicate ±1 SE. (Inset) Tracked outlines of the wall of four cells before (red) and after (blue) plasmolysis. (D) (Left) Initial images of a single cell: phase, epifluorescence, and overlay. In the overlay, the epifluorescence kymograph is false-colored green and the inverse of the phase kymograph is false-colored red. (Right) Kymographs of this cell elongating and dividing during a 400 mM hyperosmotic shock. The kymographs consist of a montage of a one-pixel-wide region around the long axis of the cell, indicated by the dashed line. The white arrow indicates the time of the shock. The red arrow indicates the period of plasmolysis during which the cytoplasm is shorter than the cell wall. The black arrow indicates the formation of a septum during cell division. (E) The concentration of sorbitol in the growth medium versus time during a hyperosmotic shock. (F) The length of the cell walls of representative cells during the same shock. The background colors indicate the four phases of response. (G) The population-averaged elongation rate of both the cytoplasm and the cell wall during the same shock (n = 42 cells). The data were smoothed with a moving-average filter with a 2-min window for illustrative purposes. The blue dotted lines indicate the steady-state elongation rates in LB (upper line) and LB+400 mM sorbitol (lower line). The confidence intervals indicate ±1 SE. The cell wall could not be tracked indefinitely owing to dilution of the cell-wall stain. (Lower) Key describing the four phases of response to hyperosmotic shock.However, bacterial cells possess several mechanisms for regulating their cytoplasmic osmolarity in response to changes in their external osmotic environment (14). Specifically, E. coli imports and synthesizes compatible solutes, and imports ions, in response to high external osmolarities (15). For example, the concentration of potassium ions in the cytoplasm scales as the external osmolality (16). Therefore, it is unclear whether raising medium osmolarity actually causes a decrease in turgor pressure over long time scales. The inverse correlation between growth rate and medium osmolarity could result from another osmotic effect such as altered water potential within the cell, which could affect, for example, protein folding (17) or signaling (18). We sought to distinguish between these possibilities by measuring the elastic strain within the cell wall as a function of medium osmolarity and by determining the time scale over which osmotic shock (acute changes in medium osmolarity) modulates growth rate and cell-wall synthesis. If growth rate scales with turgor pressure, then (i) growth rate should also scale with cell-wall elastic strain, and therefore elastic strain should decrease with increasing medium osmolarity, and (ii) growth rate, and perhaps the rate of cell-wall synthesis, should rapidly change upon osmotic shock. However, if medium osmolarity modulates growth rate independently of pressure, these processes may adapt more slowly to changes in osmolarity.We found that elastic strain decreases only moderately with increasing medium osmolarity. By measuring the growth-rate response of E. coli across time scales that spanned four orders of magnitude we concluded that osmotic shock had little effect on growth rate, except in the case of plasmolysis (when P ≤ 0, causing the cytoplasm to separate from the cell wall). The growth rate of E. coli adapted slowly to changes in medium osmolarity, over the course of tens of minutes. Furthermore, when turgor pressure was restored after slight plasmolysis, E. coli cells quickly elongated to the length that they would have attained had they never been plasmolyzed. From this result, we inferred that peptidoglycan synthesis is insensitive to changes in turgor pressure. In support of this hypothesis, we found that the speed of MreB, a protein whose motion is dependent on peptidoglycan synthesis (19–21), is largely unaffected by osmotic shock. Therefore, our results demonstrate that cell-wall biosynthesis is the rate-limiting factor in cell-wall expansion and that turgor pressure is not required for cell-wall biosynthesis, but that pressure does play a simple role whereby it stretches recently assembled, unextended peptidoglycan.     

Duplicated FoxA genes in the leech Helobdella: Insights into the evolution of direct development in clitellate annelids

Background: As an adaptation to the land, the clitellate annelid had reorganized its embryogenesis to develop “directly” without the ancestral planktonic larval stage. To study the evolution of gut development in the directly developing clitellates, we characterized the expression pattern of the conserved gut gene, FoxA, in the embryonic development of the leech. Results: The leech has three FoxA paralogs. Hau‐FoxA1 is first expressed in a subset of endoderm cells and then in the foregut and the midgut. Hau‐FoxA2 is expressed in the stomodeum, which is secondarily derived from the anterior ectoderm in the clitellates rather than the tissue around the blastopore, the ancestral site of mouth formation in Phylum Annelida. Hau‐FoxA3 is expressed during the morphogenesis of segmental ganglia from the ectodermal teloblast lineages, a clitellate‐specific trait. Hau‐FoxA1 and Hau‐FoxA2 are also expressed during the morphogenesis of the leech‐specific front sucker. Conclusions: The expression patterns suggested that Hau‐FoxA1 carries out most of the conserved function in the endoderm and gut development, while the other two duplicates appear to have evolved unique novel functions in the directly developing clitellate embryos. Therefore, neofunctionalization and co‐option of FoxA might have made a significant contribution to the evolution of direct development in Clitellata. Developmental Dynamics 247:763–778, 2018. © 2018 Wiley Periodicals, Inc.     

Direct laser manipulation reveals the mechanics of cell contacts in vivo

Cell-generated forces produce a variety of tissue movements and tissue shape changes. The cytoskeletal elements that underlie these dynamics act at cell–cell and cell–ECM contacts to apply local forces on adhesive structures. In epithelia, force imbalance at cell contacts induces cell shape changes, such as apical constriction or polarized junction remodeling, driving tissue morphogenesis. The dynamics of these processes are well-characterized; however, the mechanical basis of cell shape changes is largely unknown because of a lack of mechanical measurements in vivo. We have developed an approach combining optical tweezers with light-sheet microscopy to probe the mechanical properties of epithelial cell junctions in the early Drosophila embryo. We show that optical trapping can efficiently deform cell–cell interfaces and measure tension at cell junctions, which is on the order of 100 pN. We show that tension at cell junctions equilibrates over a few seconds, a short timescale compared with the contractile events that drive morphogenetic movements. We also show that tension increases along cell interfaces during early tissue morphogenesis and becomes anisotropic as cells intercalate during germ-band extension. By performing pull-and-release experiments, we identify time-dependent properties of junctional mechanics consistent with a simple viscoelastic model. Integrating this constitutive law into a tissue-scale model, we predict quantitatively how local deformations propagate throughout the tissue.During the development of an organism, cells change their shape and remodel their contacts to give rise to a variety of tissue shapes. Analysis of tissue kinematics has revealed that epithelial tissue morphogenesis is partly controlled by actomyosin contractility. The spatiotemporal deployment and coordination of actomyosin contractility produce shrinkage and extension of cell surfaces and interfaces, which can drive tissue invagination, tissue folding, or tissue extension (1). Understanding the mechanical nature of these processes requires force measurements in vivo; however, measurements in developing epithelia are limited, and most methods have been indirect. They rely on either force inference from image analysis (2–4) or laser dissection experiments at cell (5, 6) or tissue scales (7, 8), which provide the relative magnitude and direction of stresses from cell or tissue shape changes. In contrast, mechanical approaches have been developed in recent years to impose or measure stresses of cells in contact, including cell monolayer micromanipulation (9), pipette microaspiration on cell doublets (10), and traction force microscopy on migrating epithelia (11) and single-cell doublets (12). Recently, an elegant method using deformable cell-sized oil microdroplets has provided absolute values of stresses at the cell level in cell cultures and embryonic mesenchymes (13) but not yet in live epithelia. In this context, we sought a direct in vivo method for tension measurements and mechanical characterization at cell contacts and developed an experimental approach combining optical tweezers with light-sheet microscopy.To probe epithelial mechanics in a live organism, we chose the early epithelium of the Drosophila embryo as a model system. It consists of a simple sheet of cells that spread over the yolk and are in contact with each other through E-cadherin–based adhesion. During early embryogenesis at the blastula stage just after the end of cellularization, epithelial cells have very similar hexagonal shapes, suggesting that cell junctions have similar mechanical properties and that the internal pressure of these cells is homogeneous. At the later gastrula stage, cells undergo shape changes at distinct regions in the embryo. On the ventral side of the embryo, apical cell constriction of a few rows of cells drives tissue invagination (14), whereas on the ventrolateral side of the embryo, cell intercalation, a process whereby cells exchange neighbors by polarized remodeling of their junctions, drives tissue extension. The latter morphogenetic movement is driven by an anisotropic distribution of Myosin-II (Myo-II), which is more concentrated along junctions aligned with the dorsal/ventral (D/V) axis (15). Laser dissection of cortical actomyosin networks at cell junctions in the ventrolateral tissue has shown that such an anisotropic distribution of Myo-II causes an anisotropic cortical tension (6). However, the absolute values of tensile forces have not yet been measured, and more generally, the mechanics of cell–cell interfaces in vivo are largely unknown. Here, we addressed this issue by analyzing local mechanical measurements at cell junctions during tissue morphogenesis and determining the contribution of Myo-II to tension in this context. We determined the time-dependent response of cell–cell interfaces to forced deflection and delineated a viscoelastic model of junctions. Finally, we explored the propagation of local forces within the epithelial tissue.