Читать книгу Diatom Gliding Motility - Группа авторов - Страница 3

1 Chapter 1Figure 1.1 Drawing of a pennate diatom with two raphe branches on its valve.Figure 1.2 Hypothesis that there is a point P between apices A1 and A2, so that the apical axis is tangential to the trajectory of P.Figure 1.3 Traces of two trackers attached close to the apices of a diatom of Navicula sp.Figure 1.4 Root-mean-square deviation of the angle between the apical axis and the smoothed trajectory of the point x located between the trackers.Figure 1.5 Histogram of the frequencies of the angular difference between the direction of the diatom (apical axis) and the smoothed curve in P.Figure 1.6 Craticula cuspidata observed from an almost horizontal view.Figure 1.7 Hypothesis that there is a point P between apices A1 and A2, so that the diatom performs stochastic rotary motions around P.Figure 1.8 Root-mean-square deviation of the transverse component of the fluctuations of the hypothetical pivot point.Figure 1.9 The left side (a) illustrates the sequence of steps for reversal of direction, in which the tilting takes place after the direction of motion has been changed. In alternative (b), tilting takes place before reversing the direction.Figure 1.10 Craticula cuspidata viewed from a horizontal perspective. The transapical axis is inclined against the substrate.Figure 1.11 Trajectory of a diatom with reversal points. The point P never changes to a place of the raphe with opposite curvature.Figure 1.12 Path of a diatom of the genus Navicula with reversal points. The direction of curvature changes at each reversal point.Figure 1.13 Outline and raphe of a Cymbella.Figure 1.14 Overlay of four images from a video showing a trajectory of a diatom of the species Cymbella cistula. The yellow line shows the trajectory of the leading apex. The line segment between the apices is marked in white.Figure 1.15 On the left (a) the superimposition of images of a Cymbella rotating around a point near the helictoglossa is shown, on the right (b) a sketch of the diatom with raphe.Figure 1.16 Surirella biseriata in valve view.Figure 1.17 Trajectory of a Surirella biseriata. The driving side changes at the reversal points.Figure 1.18 Paths of Surirella biseriata in a culture. They were visualized by overlaying frames of a video.Figure 1.19 Pinnularia viridiformis with a length of approx. 90 μm.Figure 1.20 Places within and on a biofilm where Pinnularia viridiformis can be found. The typical movement patterns are indicated by arrows. Shunting movements are marked with short arrows at both apices.Figure 1.21 Superimposed frames of a video during the standstill of a diatom. A tension has built up in the biofilm.Figure 1.22 Nitzschia sigmoidea on the water surface viewed with PlasDIC.Figure 1.23 Nitzschia sigmoidea with a stereomicroscope in oblique view.Figure 1.24 Sketch of a Nitzschia sigmoidea on the water surface seen from the horizontal direction.Figure 1.25 Sketch of two adjacent Nitzschia sigmoidea on the water surface seen from the horizontal direction.Figure 1.26 Very regular structure of a diatom cluster on the water surface (dark field).Figure 1.27 Relative speed of two diatoms plotted versus their distance.Figure 1.28 Energetically favorable patterns of three diatoms on the water surface: all diatoms parallel (a) and diatoms form a triangle (b).Figure 1.29 Frequently observed movement patterns: movement along the raphe (a) and angular changes at connected apices (b).Figure 1.30 Image sequence showing the temporal development of seven connected diatoms. The time between the first and last image is 170 seconds.Figure 1.31 Pinnularia gentilis.Figure 1.32 Cymbella lanceolata.Figure 1.33 Two small colonies photographed with PlasDIC.Figure 1.34 Elementary steps that contribute to structure formation.Figure 1.35 Movement activity of diatoms between colonies.Figure 1.36 Colonies at the beginning of intensive light irradiation (a) and after about two hours (b).Figure 1.37 Cymbella culture in the light phase (a) and dark phase (b).Figure 1.38 Number of free diatoms (blue) and the number of diatoms bound in colonies (red) over 24 days. A yellow bar indicates the phases of bright light.Figure 1.39 Total number of diatoms (red) with exponential fitting (blue).Figure 1.40 Number of diatoms in motion over the last 10 observation days.

2 Chapter 2Figure 2.1 The polycarbonate channel used to image diatom motion, photographed on the stage of the inverted microscope. The depth, width, and length of the channel are 1 mm, 2 mm and 60 mm, respectively. A glass slide was bound to 2.5 mm × 7.5 mm polycarbonate block using two pieces of double-sided tape placed on the polycarbonate block. The coordinate origin coincides with the origin for the pixels. Diatoms move on the glass slide at the bottom of the chamber. The inset figure shows a schematic diatom, the coordinate axes, and the origin, in the plane of the top of the slide.Figure 2.2 (a-b) Scatter plots of the simulated test object and the stationary polystyrene particle centroids, respectively. The stationary polystyrene particle was imaged at 821 fps. The number of points plotted is 1000 for the simulated particle and 4000 for the polystyrene particle.Figure 2.3 Displacement histogram of the diatoms (symbols) and stationary polystyrene particle (continuous line) imaged at 821 fps.Figure 2.4 (a) Plot of total displacement as a function of time in a diatom centroid measurement. The frames around the arrow are investigated in detail. (b) Overlay image of the diatom boundary for frames where the displacement was 293 nm at 1986 ms. White pixels show the overlapped points in the diatom boundary, while colors indicate change in boundary location; (c) and (d) show two consecutive frames of the diatom (diatom #2) for the displacement shown by an arrow in Figure 2.4 (a). The boundaries were calculated using the “Binary Centroid” algorithm. In b, green = frame c and magenta = frame d.Figure 2.5 Angular velocity histogram of the diatoms #1-3 that were imaged at 821 fps. The diatoms exhibited changes in their orientation angles as they traversed the imaging strip horizontally.Figure 2.6 In Figures 2.6a and 2.6c the orientation angle corresponding to the diatoms whose trajectories were previously investigated are shown (diatoms #2 and #3 in a and c, respectively). In Figure 2.6e the orientation angle of another diatom (diatom #4) is shown. Orientation angle is the angle between the major axis of the diatom and the x-axis. Figures (b), (d), and (f) are overlay images of the diatom boundary for different frames of the data shown in (a), (c), and (e), respectively. White pixels show the stationary points in diatom boundary, while colors indicate change in boundary location. An extra Gaussian smoothing was included in the image processing algorithm for the diatom #4, whose orientation angle is shown in Figure 2.6e. Otherwise, it was not possible to extract orientation data for diatom #4.Figure 2.7 (a) The diatom in Figure 2.6d is rotated around the arc center of the white pixel region in Figure 2.6d; (b) the consecutive intersections of the major axis of the diatoms during rotation are shown as yellow dots. In (b), while the green dots at the centerlines are close to one another, they would coincide exactly if the rotation were precisely around the center of the diatom.Figure A1 Displacement histograms of the image of the stationary polystyrene particle situated in the micromachined channel; (a) and (b) show x and y displacement histograms, respectively. Dots represent microscopy measurement while continuous lines are the Gaussian fits to the measurements. Two Gaussian terms are used for fitting total displacement data. The particle was imaged at 821 fps.Figure A2 Fast Fourier Transform (FFT) of the x and y coordinates of the centroid location for the recordings made with the stationary polystyrene particle and a diatom; (a) and (c) are the FFTs of the x centroid data and (b) and (d) are the FFTs of y centroid data of the polystyrene particle and diatom #3, respectively. Prior to the FFT the centroid location mean was subtracted from the data.Figure A3 Representative trajectories of the centroids of diatoms #1, #2, and #3 are given in (a), (b), and (c), respectively, over 500 frames. The diatoms were imaged at 821 fps.Figure A4 Zoomed in views of the large displacements in the representative trajectories given in Figure A3. The plots (a), (b), and (c) show the centroids of diatoms #1, #2, and #3, respectively.Figure A5 Mean square displacement (MSD) data of diatom trajectories. Diatom motion recordings were divided into segments that are 500 frames long and MSD data of each segment was calculated. The black dashed line represents the mean MSD and red line represents the best fit to the MSD for all diatoms (diatom #1 – (a), diatom #2 – (b), diatom #3 – (c)). All MSD data falls into the range shown by gray shaded areas in the figures.Figure A6 Normalized velocity autocorrelation of all the diatoms (diatom #1 – (a), diatom #2 – (b), diatom #3 – (c)). The same trajectory segments as in calculation of diatom MSD data were used. All repetitions were used in calculating the autocorrelation.Figure A7 The publicly available video [2.87] was analyzed using the SURF feature detection algorithm (please see the main text for the details); (a) displacement calculated in pixels as a function of the frame number; (b) the overlay of the features detected at frame numbers 422 (crosses) and 423 (circles). pts = points. Scale bar was not present in the original video.

3 Chapter 3Figure 3.1 Vesicles carrying mucilage (black arrows) on sections of cells of Pleurosigma sp. near the girdle bands (a) and Encyonema ventricosum (b) in the area of areolae (TEM). Around the cells Encyonema ventricosum are visible fibers between the plasmalemma and the frustule and a dense layer of mucilage fibers on the surface of the frustule, however, the connections between them are absent. Chl - chloroplasts; m - mitochondrion. Scale – 200 nm.Figure 3.2 The surface cells of Nitzschia sp. (a, b) and Pleurosigma sp. (c, d) after removal of frustules (SEM). At the tips of the precisely repeated contour of the frustules in the area of the raphe system of Nitzschia sp. in some cases separated fragments (arrows) are visible; it may be the place where the cell is firmly attached to the valve. The frustule of Pleurosigma does not have so extremely relief, but the contours of the cell here is very accurately repeated especially in the area of the raphe (arrows). It is possibly that most of the surface is not diatotepum, as this polysaccharide layer is very firmly attached to the leaf, which is very well illustrated by the E. ventricosum (Figure 3.1b). Scale: (a, d) – 10 μm; (b) – 5 μm; (c) – 1 μm.Figure 3.3 Secretion of mucilage (arrow) through the raphe of the E. ventricosum (a) and vesicle (arrow), containing mucus near the raphe of Pleurosigma sp. (b) on ultra-thin cross sections (TEM) Scale: (a) – 200 nm; (b) – 500 nm.Figure 3.4 Staining of actin microfilaments (phalloidin Alexa Fluor 488, green fluorescence) and nuclei (DAPI, blue fluorescence) in Pleurosigma sp. (a-b’) and Nitzschia sp. (c, d). Scale: (a, b) – 15 μm; (b’) – 5 μm; (c, d) – 10 μm.

4 Chapter 4Figure 4.1 Morphology of an araphid diatom (a), Staurosira construens var. venter (scale bar: 2 μm) and a raphid diatom (b), Navicula radiosa (scale bar: 10 μm), diatom on valve view. The raphe (indicated by a white arrow) runs through the whole valve and is a primary structure for adhesion to surfaces and moving. (c) A close-up of the raphe (indicated by a white arrow) is shown (scale bar: 1 μm). (SEM images downloaded from diatoms.org with permission [4.107] [4.120] [4.143].)Figure 4.2 Schematic of a benthic biofilm. Algae, predominantly pennate biofilms together with bacteria, protists, and fungi, are embedded within a protective matrix of extracellular polymeric substances (EPS). A benthic biofilm experiences high variability of spatial and temporal gradients of environmental factors such as inorganic nutrients, dissolved organic matter (DOM), and light. (Figure from Sabater et al. [4.129]. Reprinted under CC-BY license.)Figure 4.3 A summary of some factors and gradients affecting diatom adhesion and motility on intertidal sediment. While adhering to the surface, cells sense their wettability through the production of intracellular nitric oxide (NO), which in turn mediates EPS production. Pennates generally have a preference for hydrophobic surfaces for attachment. Shear force or water flow affects substrate attachment with weakly attached cells easily dislodged by stronger shear. Once attached, the gliding capability and reversal control are mediated by the availability of extracellular and intracellular Ca²⁺, respectively. As light only penetrates a few mm on the sediment, the substrate is divided into the photic and aphotic zone. Cells undergo diel vertical migration to the photic zone to photosynthesize on the surface with a taxa-specific temporal rhythm, thus leading to micro-niche segregation. Nutrient concentrations vary with depth, and motile raphids take up dissolved silicate (dSi) and phosphate (dP) on the aphotic zone as there are higher concentrations of these mineral nutrients with depth. The aphotic zone also provides a protected and stable environment for vegetative and sexual reproduction. (Figure modified and redrawn from Saburova et al. [4.132] and adapted from Fenchel [4.59]. Reprinted with permission.)Figure 4.4 (a) Scanning electron microscopy (SEM) images of EPS or mucilage trails and strands from the motile raphid diatom, Navicula sp. (scale bar: 5 μm). EPS trails were observed to be either straight or curved. (SEM images from Chen et al. [4.26]. (Reprinted under CC-BY license); (b) Adhesion of Nitzschia palea towards surfaces with different surface wettabilities (scale bar: 15 μm). Cells adhered more to hydrophobic surfaces after 4 hours. Insets are duplicate experiments. (Optical microscopy images from Laviale et al. Reprinted (adapted) with permission from Laviale et al. Copyright 2019 American Chemical Society [4.91]).Figure 4.5 (a) A representative cell trajectory of Seminavis robusta showing the run-reverse motility of cells from high cell density experiments. (Cell track replotted from data from Bondoc et al. [4.15]). (b) The circular run-reverse gliding technique observed in Nitzschia communis under isotropic environmental conditions. Cells form arc-like runs with constant speed, reverse, and continue the arc-like runs in the opposite clock direction. Filled squares signifies the starting point of the cells. (Figure from Gutiérrez-Medina et al. [4.68]. Reprinted with permission.)Figure 4.6 Motility of the model raphid pennate Seminavis robusta towards different stimuli across its life cycle. For every mitotic division, cells undergo size reduction until they reach a sexual size threshold (SST). Cells can either continue to undergo vegetative growth or sexual reproduction to reconstitute their size and escape death. Throughout vegetative growth, cells require nutrients. Gradients of dissolved silicate (dSi) and phosphate (dP) elicited starved cells to accumulate at point sources within 5 and 20 min, respectively. Dissolved germanium (dGe) did not elicit any attraction, pointing to substrate specificity response. Meanwhile, starved cells also did not respond to gradients of dissolved nitrate or ammonium (collectively called dN). Once cells reach SST, they release sex-inducing pheromones (SIPs) that control the production of diproline (DPR) on MT^– cells. This pheromone is used by the MT⁺ cells as a chemical guide on locating MT– spatially and pair with it. Sexual reproduction requires trace amounts of dSi for the reconstitution of the silica frustule of the initial cells. Additionally, SIP priming is essential for cells that recently crossed SST. On the other hand, critically small-sized cells can bypass the priming process and be readily attracted to diproline. This self-priming could be a self-preserving strategy for the cells to avoid extinction. (Figure from Bondoc et al. [4.16]. Reprinted with CC-BY license.)

5 Chapter 5Figure 5.1 The effect of incubation time on Stauroneis response times in a mixed culture of Craticula cuspidata and Stauroneis phoenicenteron. This graph displays the average direction change response times for Stauroneis phoenicenteron cells in the presence of ca. 9:1 ratio of live C. cuspidata: S. phoenicenteron cells. S. phoenicenteron cells were isolated and washed from culture and incubated together with C. cuspidata cells (C/S) in a 9:1 C. cuspidata:S. phoenicenteron ratio. Cells were then irradiated at their leading end with high irradiance (ca. 10⁴ μmol/m²s) 1s pulses of blue (470 nm) light, and observed to determine the time until they changed direction (response time). Response times significantly increased almost 2-fold from the initial incubation interval (0-10 min) within 20-30 min (30±2 μm/s and 57±7 μm/s respectively, P=0.003). Graphs represent the mean values of response times ± 1 SE. For comparison, unirradiated Stauroneis cell response times were 155±11 μm/s. Error bars represent ± 1 SE.Figure 5.2 The effect of Craticula cuspidata or Pinnularia viridis presence on Stauroneis phoenicenteron response times. This graph displays the average direction change response times for S. phoenicenteron cells alone on slide chamber (Control). On a mixed slide chamber in a group by themselves (Isolated) or in a slide chamber in the presence of a high ratio of live C. cuspidata or P. viridis cells to S. phoenicenteron cells. Cells were then irradiated at their leading end with high irradiance 1s pulses of blue light, and observed to determine the time until they changed direction (response time). The presence of either C. cuspidata or P. viridis caused significant increases (P < 0.03) in cell response time from either control or isolated S. phoenicenteron. Error bars represent ± 1 SE.Figure 5.3 The effect of culture medium on Stauroneis phoenicenteron response times. This graph displays the average direction change response times for S. phoenicenteron cells placed in the presence of culture medium in which either S. phoenicenteron, Craticula cuspidata, or a mixture of S. phoenicenteron and C. cuspidata cells had been growing. Samples of the media were extracted, then briefly centrifuged in a microcentrifuge to remove any contaminating cells or cell debris. The S. phoenicenteron cells were then immersed into the desired medium, then isolated onto a slide chamber, allowed to incubate, then irradiated at their leading end with high irradiance 1s pulses of blue light, and observed to determine the time until they changed direction (response time). There was no significant difference between any of the treatment groups. Error bars represent ± 1 SE.Figure 5.4 The effect of Pinnularia viridis on Stauroneis phoenicenteron response times. This graph displays the average direction change response times for S. phoenicenteron cells placed in two different areas of a cell chamber with differing proximities to P. viridis. This graph displays the average direction change response times for unirradiated S. phoenicenteron (Unirradiated), S. phoenicenteron irradiated in a slide chamber by themselves (Control), and S. phoenicenteron placed in a two-area slide chamber in which the entire chamber was subject to the same medium. Cells were irradiated at their leading end with high irradiance 1s pulses of blue light, and observed to determine the time until they changed direction (response time). The S. phoenicenteron in the two-area chamber were either in an area by themselves (Isolated), or in an area of the chamber where they were in close proximity to a large number of P. viridis (Live Pinn). In some trials the S. phoenicenteron were placed in the presence of dead P. viridis (Dead Pinn) that had been killed by immersing the P. viridis cells for 30 sec in 95% ethanol prior to rinsing the P. viridis with distilled water and fresh diatom medium. The S. phoenicenteron cells in the presence of living P. viridis showed a significant increase in response time, even though they were exposed to the same medium as the isolated S. phoenicenteron. while those in the presence of dead P. viridis showed no such increase in response time. Error bars represent ± 1 SE.Figure 5.5 Effect of Craticula cuspidata fixation method on repression of Stauroneis phoenicenteron direction change response time. S. phoenicenteron cells isolated and washed from culture were incubated together with C. cuspidata cells in a VALAP-spaced cell chamber where some Stauroneis were in contact with Craticula (grey bars) while other Stauroneis (isolated, solid bars) had no Craticula in near proximity. Stauroneis cells were then irradiated at their leading end with high irradiance flashes of blue light, and observed to determine the time until they changed direction (response time). Stauroneis in a mixed region with live Craticula had a significantly greater response time than Stauroneis in an isolated area (P = 0.01). This increase in response time remained when S. phoenicenteron were in the presence of C. cuspidata cells killed in ethanol or acetone fixation alone, but no significant difference in response time was observed when C. cuspidata were killed with a 50:50 acetone:ethanol mixture. The isolated S. phoenicenteron in the chamber also showed increased response times in the treatments containing ethanol or acetone fixed C. cuspidata. Graphs represent the mean values of response times ± 1 SE.Figure 5.6 Histogram analysis of Stauroneis phoenicenteron response times. The figure displays the distribution of direction change response times for S. phoenicenteron when in isolated groupings (a) or when in a mixed assemblage with Pinnularia viridis (b). Stauroneis phoenicenteron cells, when in the presence of P. viridis show some cells with control levels of rapid response times, along with a number of cells with greatly increased response times.

6 Chapter 6Figure 6.1 Low-temperature scanning electron micrographs of diatom life styles (clockwise from top left): Stalked, tube forming, adpressed, epipelic. (Images: Irvine Davidson, University of St Andrews.)Figure 6.2 Low temperature electron micrograph of the pathway of an epipelic diatom moving thorough fine silt (Bar marker = 10 um). (Image: Irvine Davidson, University of St Andrews.)Figure 6.3 Top: Change in environmental driver (pressure) dependent on the presence of a “stabilizing biofilm” or under “regular erosion.” Bottom: Low-temperature scanning electron micrograph of MPB biofilm structure. (Image: Irvine Davidson, University of St Andrews.)

7 Chapter 7Figure 7.1 Proposed model of the control of vertical migration by sediment-inhabiting benthic pennate diatoms, as responding to main directional environmental stimulus, light and gravity. The figure illustrates the variation with the time of day of the diatom biomass at the sediment surface on samples kept in the dark (closed circles) and exposed to constant low light (150 μmol m-² s-¹) during the subjective low tide period (open circles) (for more details, see [7.18]). Example of a day when the low tide takes place during the middle of the day. The gray horizontal plots represent the strength (bar thickness) of photo- and geotaxis, of negative and positive signal, along the day. (1) Upward migration starts before the beginning of the light period, driven by negative geotaxis. (2) Negative geotaxis ceases roughly at the time expected for start of the low tide light period: if no light is available at surface, the diatoms stop migrating upwards and the incipient biofilms start to disaggregate, due to random cell movement or weak positive geotaxis. (3) During light exposure, cell movements are controlled mainly by phototaxis, either positive (under low light intensities) or negative (under high light intensities). In the particular case of the data in the figure, positive phototaxis dominates, as samples were exposed to low intensity. (4) Anticipating the end of the light period, geotaxis becomes dominant over phototaxis, as cells begin to migrate downwards without any changes in incident illumination. Vertical gray areas represent periods of darkness. Vertical white area represents the period of light exposure (150 μmol m-² s-¹) of the light exposed samples.

8 Chapter 8Figures 8.1–8.13 LM images of Eunotia taxa in valve view (Figures 8.1–8.8) and girdle view, ventral-side up (Figures 8.9–8.11) to show variation in morphology and raphe shape and location (arrow in some images). (Figure 8.1 [8.28]) E. bilunaris (Ehrenb.) Souza – raphe recurved almost 180°. (Figure 8.2) E. serra Ehrenb. – raphe on valve face with slight curve toward apex. (Figure 8.3 [8.28]) E. areniverma Furey, Lowe et Johansen – raphe follows margin of apex from ventral to dorsal margin, and up onto dorsal mantle. (Figure 8.4 [8.28]). E. pectinalis var. ventricosa (Ehrenb.) Grunow (E. pectinalis var. ventralis (Ehrenb.) Hust. = synonym). (Figure 8.5) E. bigibba Kütz. – raphe with slight recurve. (Figure 8.6) E. incisa Smith ex. Gregory – raphe (not visible) only present on valve mantle (see Figure 8.17). (Figure 8.7) E. muscicola Krasske – raphe (not visible) on the valve face with slight recurve (see Figure 8.20). (Figure 8.8) Close up of valve apex and raphe of E. bilunaris in Figure 8.1. (Figure 8.9) E. bigibba and (Figure 8.10) E. tetraodon Ehrenb. – ventral-girdle view. Depth of valve only permits part of raphe to be in focus (on one plane) at a time. (Figure 8.11) Unknown valve in ventral-girdle view. Raphe almost all in focus. (Figure 8.12) Epiphytic cells of E. bilunaris (E) standing up on end from the mucilaginous sheath(s) of the cyanobacterium Hapalosiphon Nägeli ex Bornet et Flahault (Image credit R.L. Lowe). (Figure 8.13) SEM image of Eunotia on a bryophyte. For all images except for Figure 8.8: black scale bar = 10 μm. Figure 8.8: white scale bar = 5 μm. (Figures 8.1, 8.3, 8.4 originally published in Furey et al. [8.28] www.schweizerbart.de/journals/bibl_diatom).Figures 8.14–8.22 SEM images of Eunotia taxa to show variation in morphology, along with external and internal raphe (R) shape and location on the valve face and valve mantle, helictoglossa (h), shape, location, and internal expression of the rimoportula (r), and external expression of the rimoportula pore (rp). (Figure 8.14 [8.28]) E. areniverma – raphe follows margin of apex from ventral to dorsal margin, and up onto dorsal mantle. (Figure 8.15 [8.28]) E. areniverma – internal view of apex. Rimoportula located mid apex. (Figure 8.16 [8.28]) E. pectinalis var. ventricosa – external view showing path of raphe from mantle onto valve face with slight recurve. (Figure 8.17 [8.28]) E. incisca – raphe located completely on valve mantle. External expression of rimoportula. (Figure 8.18) E. bigibba – curve of raphe onto valve face with slight recurve. (Figure 8.19) E. serra – raphe on valve face with slight curve toward apex. (Figure 8.20) E. muscicola – curve of raphe onto valve face with a slight recurve and (Figure 8.21) internal view of valve apex with rimoportula located close to the helictoglossa. (Figure 8.22) E. serra – internal view of valve apex with rimoportula located closer to dorsal margin. Scale bars as shown. (Figures 8.14–8.17 originally published in Furey et al. [8.28] www.schweizerbart.de/journals/bibl_diatom). [8.38] [8.53] [8.66], though their position could be derived if a more complex raphe system became reduced (discussed by Kociolek [8.44] and Siver and Wolfe [8.69]). Movement in eunotioid diatoms with their short raphe system, typically described as slightly (< 2 µm/sec [8.31]) or weakly motile (2 to 4 µm/sec [8.31]) (see eunotioid taxa in DONA [8.21] – Furey [8.26]), contrasts with that of more motile forms like naviculoid, nitzschioid, or surirelioid diatoms with more extensive raphe systems, typically described as moderately to highly motile. Examination of motility in Eunotia species may provide unique insight into motility in diatoms, especially for diatoms with more complex raphe systems. A search for the terms “motile” or “move” in florae focused on Eunotia (e.g., [8.20] [8.28] [8.48] [8.51]) and in >40 manuscripts with descriptions of Eunotia taxa new to science revealed little to no mention of motility. This chapter focuses on motility in the diatom genus Eunotia, but does not cover cellular or biomechanical details around the mechanisms of movement, as other chapters in this book discuss these aspects at length.Figures 8.23–8.27 Schematic representation of some of the movement types for Eunotia. (Figure 8.23) Schematic of forward movement – apical displacement where cells tilt slightly so the anterior ends of the valves remain in contact with the surface and the posterior ends become slightly raised (**schematic modeled after Palmer [8.60] plate vi. fig. 2, and Bertrand [8.6] fig. 1**). (Figure 8.24) Valve in girdle view. (Each raphe branch labeled after Bertrand [8.6] (**see also Harbich [8.30]**). Black arrow following the line of the raphes on B to C apices represents diagonal line of direction, where the raphe on the C end becomes active. (Figure 8.25) Black horizontal arrow represents diagonal line of direction. Bidirectional arrow shows transition between raphes involved in forward motion (**schematic modeled after Bertrand [8.6], fig. 1**). (Figure 8.26) Schematic of a vertical polar pivot which can return a cell in girdle view, dorsal-side down (a) to ventral-side down (b,c) where a cell can then continue forward movement (c) (**schematic modeled after Palmer [8.60], plate vi. fig. 2, and Bertrand ([8.6], fig. 5]**). (Figure 8.27) Schematic of a horizontal, polar pivot (a,b) to show direction of raphe activity (straight arrows, A and B). Note the cell is depicted ventral side up so the raphe branches are visible (rather than dorsal side up as the movement occurs). Curved arrows show direction of rotation. Dot represents the pivot point. (**Modified after images from Harbich [8.30]). See additional schematics in Bertand [8.5].

9 Chapter 9Figure 9.1 Gliding cells of Nitzschia sigmoidea with stalked epiphytes of Pseudostaurosira parasitica: a single epiphyte in connective view (a), in valve view (b), and two epiphytes attached to the same frustule (c), which can be seen in movement [9.39].Figure 9.2 Cells of Nitzschia sigmoidea with adnate epiphytes of Fallacia helensis (OM): single epiphyte in connective view and host in valve view (a), four epiphytes in valve view and host in connective view (b), which can be seen in movement [9.43].Figure 9.3 Cells of Nitzschia sigmoidea with many epiphytes (OM): epiphytes of Amphora copulata on a still gliding host (a), epiphytes of Amphora copulata on a dividing host (b), epiphytes of Amphora copulata and Pseudostaurosira parasitica on the same host (c), which can be seen in movement [9.41].Figure 9.4 Focus on epiphytes of Nitzschia sigmoidea (SEM). Adnate, Fallacia helensis, and stalked, Pseudostaurosira parasitica, epiphytes on the same host (a), two cells of Pseudostaurosira parasitica still associated after division (b), two superimposed cells of Fallacia helensis after division and a third single one attached on the edge of the frustule (c), apex of a cell of Pseudostaurosira parasitica with the mucilaginous pad secreted for adhesion (d), individual of Amphora copulata (e) and internal view of one valve of Amphora sp., probably Amphora copulata var. epiphytica Round & Kyung Lee, considering the almost circular areolae on the ventral side (f). Scale bars indicate 10 µm, except in 9.4d (=5 µm).Figure 9.5 Variations in the specific composition of epiphytes on Nitzschia sigmoidea between two sampling sites located on two connected rivers (up- and downstream sites), expressed as the occurrence of three epidiatomic species on frustules of N. sigmoidea. In fact, through the observation of fresh material, N. sigmoidea could not be strictly distinguished from the close species N. vermicularis. Both species were present in each site (see [9.44] for details).Figure 9.6 Sigmoid frustules of Nitzschia sigmoidea (NSIO) and Gyrosigma attenuatum (GYAT) (OM, H₂O₂ treated material). Two species co-occurring in rivers samples with similar abundance, valve length and motility. However, Gyrosigma attenuatum was never seen with epiphytes.

10 Chapter 10Figure 10.1 Drawing adapted from O.F. Müller (1783, translated in [10.54]), who was the first to characterize Bacillaria colonies. Examples 1 through 8 show the various states of expansion and contraction (dynamic phenotypes) of colonies.Figure 10.2 Bacillaria close-up images of single cells using scanning electron microscopy (SEM). (a) a whole valve seen from the inside. (b) close up of the same, middle section. The horizontal slit is the raphe. It lacks a central node. (c) Tip of the inside of a valve. (d) Middle section of a valve, exterior view. Note the raphe is a slit through the whole valve. (e): External view of the tip of a valve [10.33]. (Reprinted with permission of Amgueddfa Cymru, National Museum Wales.)Figure 10.3 Demonstration of the image tracking procedure. (a) definition of tracked feature (white ellipse within a cell). (b) labeled (numbered) cells with relative measurements provided in red. The determined coordinates refer to a target in the middle of the template. The target can be moved and placed on the apex of the diatom being tracked. Then the coordinates of the apex are captured. This position is indicated in Figure 10.3a by a mark and a vertical line. Image scale: 38.36 μm per cm, or 0.325 μm per pixel. Scale bars 50 µm.Figure 10.4 A diagram showing the five points on a sample cell (two ends, midpoint of the transverse line, and edges of the cell).Figure 10.5 Point A is on the edge (vertical direction). The gradient direction is normal to the edge. Points B and C are in gradient directions. So, point A is checked with point B and C to see if it forms a local maximum. If so, it is considered for the next stage, otherwise, it is suppressed (set to zero). The result is a binary image with “thin edges.”Figure 10.6 Diagram showing an example of hysteresis thresholding and the labeled edge relative to the “sure edge” threshold (V_max).Figure 10.7 An example of feature identification performance for the Watershed Segmentation algorithm (left, red boundary) and Canny Edge Detection algorithm (right, white boundary). Image scale: 38.36 μm per cm, or 0.325 μm per pixel. Scale bars 50 µm.Figure 10.8 An example of feature identification training (purple rectangles) for the deep learning approach on a single set of cells. Notice the resolution of the colony. An example of correct performance is shown in Figure 10.5. Image scale: 38.36 μm per cm, or 0.325 μm per pixel. Scale bar 50 µm.Figure 10.9 Rank-order analysis of bounding box (cell) sizes (area) across the dataset. The area is measured in pixels squared. Image scale: 38.36 μm per cm, or 0.325 μm per pixel.Figure 10.10 Rank-order analysis of height (blue) and width (red) of bounding boxes (cells) across the dataset. The area is measured in pixels squared. Image scale: 38.36 μm per cm, or 0.325 μm per pixel.Figure 10.11 (Top) location of centroids in normalized coordinate space in selected dataset for static analysis. (Bottom) First two principal components from PCA analysis (PC1 represents horizontal position, while PC2 represents vertical position) of coordinates representing the x,y position for all four edges of each bounding box using the selected datasets for static analysis. Image scale: 38.36 μm per cm, or 0.325 μm per pixel.Figure 10.12 An example of feature identification optimization procedures implemented in DeepLabv3. GRAY: no optimization applied, RED: Optimization #1, BLUE: Optimization #2. Given an initial number of training frames (y-axis), the non-optimized procedure (originally detected) will yield a certain number of boxes (x-axis). Applying various optimization procedures generally leads to a decreased number of boxes per frame for both low and high numbers of boxes.Figure 10.13 Four examples of how the identified features map to two different images (a, c, e, g) of a Bacillaria colony. Points (b, d, f, h) represent the centroids for all bounding boxes identified in images a, c, e, and g, respectively. Image scale: 38.36 μm per cm, or 0.325 μm per pixel. Image scale: 38.36 μm per cm, or 0.325 μm per pixel. Scale bars 50 µm.Figure 10.14 Three examples of how images of a Bacillaria colony are converted into a skeleton image. (Top Row) light microscopy images, (Middle Row) thin skeletonization based on a procedure implemented in GIMP, (Bottom Row) thick skeleton based on a procedure implemented in GIMP. Image scale: 38.36 μm per cm, or 0.325 μm per pixel. Image scale: 38.36 μm per cm, or 0.325 μm per pixel. Scale bars 50 µm.Figure 10.15 Examples of relative movement of cells in a sample colony. (a) Comparisons between changes of position for cell #2 relative to cell #1 (red) and cell #3 relative to cell #2 (blue). (b) Comparisons between changes of position (red) and changes of velocity (blue) for cell #2 relative to cell #1. (c) a phase diagram of the data shown in b, velocity versus position. (d) comparison between changes of position for cell #2 relative to cell #1 (red) and sine wave (black dashed). The oscillation period in frame c is 62.56 seconds.

11 Chapter 11Figure 11.1 Expansion of a cylindrical mucopolysaccharide fibril.Figure 11.2 Schematic drawing of the expanding fibril.Figure 11.3 Time dependence of the sound pressure p.Figure 11.4 Time dependence of the sound pressure p for different mass transfer coefficients k.Figure 11.5 Schematic drawing of a jar with diatoms.Figure 11.6 Raphid diatom (to our best knowledge), caught by FZ; Note the visibility of the chloroplasts inside the diatoms; the scale bars are estimated.Figure 11.7 Stones from underwater with golden-brown film on them, collected by FZ in Natschbach, Austria, on May 5, 2019.Figure 11.8 Comparison of diatom samples obtained with different methods; the scalebars are estimated.Figure 11.9 Setup of first measurements.Figure 11.10 Measurement of jar with diatoms (right), reference jar (left).Figure 11.11 Jars with diatoms and little glass balls.Figure 11.12 Spectrogram of measurement with diatoms.Figure 11.13 Averaged spectrum over the whole spectrogram (0.5 s – 3.5 s).Figure 11.14 Averaged spectrum over the range 2.75 s – 3 s.Figure 11.15 Averaged spectrum between 1 s – 1.5 s.Figure 11.16 Spectrogram of reference measurement.Figure 11.17 Averaged spectrum over whole spectrogram (0.5 s – 3.5 s).Figure 11.18 Averaged spectrum between 0.5 s – 0.65 s.Figure 11.19 Averaged spectrum between 1.85 s – 2 s.Figure 11.20 Averaged spectrum between 2.5 s – 3.0 s.

12 Chapter 12Figure 12.1 Waves formed by the microfibrils. The waves for each half­raphe vary independently in frequency and intensity. Translation of labels: Sens de progression l’onde = Direction of wave progression; Chloroplastes = Chloroplasts, Noyau = Node; Diatomée en vue connective = Girdle view of diatom.Figure 12.2 Movements of the microfibrils after their shortening: Position 1: Microfibrils at rest; Position 2: rc, shortening of one of the two microfibrils; Position 3: The two microfibrils, bonded at the point of connection, slanted at an angle according to the value of the shortening. Translation of labels: Sens du mouvement des microfibrilles = Direction of microfibril movement; Point de liaison = Contact point; Base des microfibrilles = Microfibril base. Top: Variation of f as a function of rc. Bottom: General formula f = (e2 – rc2 + 2L. rc) / 2rc. Setting L = 1, f = (e2 – rc2 + 2rc)/2rc. Setting L = 1, e = rc, then f = 1 = L = 90°.Figure 12.3 The slope of myosin heads generates the sliding of the actin filaments which causes their shortening and the retraction of the microfibrils. Translation of labels: Phase de contraction des microfibrilles (raccourcissement relatif) = Microfibril contraction phase (relative shortening); Phase de repos = Resting phase; Génération d’impulsions par dépolarisation des charges électriques au niveau du plasmalème = Depolarization pulse generation by electric charges at the cell membrane level.Figure 12.4 Birth, growth and slope of the microfibrils in relation to the wave trains of contraction. At rest, there is expulsion of mucus. Translation of labels: Inclinaison et “coup de fouet” des microfibrilles = Tilt and “whiplash” of microfibrils; Expulsion du mucus = Expulsion of mucus; Redressement des microfibrilles = Recovery of microfibrils; Repos = Rest; Diatomée = Diatom; Train d’ondes de contraction = Contraction of wave train.Figure 12.5 Displacement of the diatom on a half­raphe with organic matter drive on another half­raphe. Translation of labels: Émission de mucus = Mucus emission; Matière organique entraînée = Attached organic material; Sens de progression l’onde = Direction of wave travel; Résidue de mucus = Residue mucus²; Sens du mouvement de la diatom = Direction of diatom movement.

13 Chapter 13Figure 13.1 Appearance of Navicula sp. diatoms under an optical microscope. Cells are about 14 µm long [13.24].¹Figure 13.2 SEM Image of Navicula sp. in girdle band face (a) and the frustule view (b).²Figure 13.3 Pore array and pore structure on the frustule. (a-c) Meso-porous structure in the frustule, (d) nanoporous strucuture inside mesopores. AFM mapping of pores around ridge (e) and mesopores (f).³Figure 13.4 The bending ability of the diatom wall during the re-positioning process, scale bar is 5 μm. (a-c) The diatom indicated by the arrow attaches to the wall and the diatom on the right side is changing its orientation by rotating in situ, which the maximum bending angle is about 37 degrees in (b).Figure 13.5 Equipment for analyzing bending ability of diatoms. See text for description. (a) Experimental set up of electrode corrosion for preparing tungsten needles, (b) characterization of an as-prepared tungsten needle, (c) experiment set up of bending ability characterization inside the SEM equipped with a micromanipulator and a force sensor.⁴Figure 13.6 The relationship between bending deformation and stress of frustules. The scale bar is 5 μm. (a) original position, (b) bending deformation with an angle of 26 degree, (c) forces measured with deformation.Figure 13.7 Simplifying the frustule bending into a simple cantilever beam system.Figure 13.8 Classical Edgar model. See text for description of components.Figure 13.9 Pits found in the mucilage trails. Scale bar equals 2 μm.⁵Figure 13.10 Diatoms locomoting while raised at an angle of inclination.⁷Figure 13.11 Cross-section view of diatom locomotion, (a) normal locomotion, (b) inclined locomotion.Figure 13.12 The same diatom crawls along the raphe (a-b) and its girdle band (c-f) surfaces.⁸Figure 13.13 Diatoms crawl with the girdle band facing the substrate.Figure 13.14 The circular structures in the body of some locomoting diatom cells. (a-d) are serial captures of the same diatom while locomoting, which two circular structures were observed to move within the cell body at high-frequency vibration at the microscale.⁹Figure 13.15 Obscure circular structures in locomoting diatom cells. Circular structures could be observed in some diatoms (a) but not always could be found (b) even under LSCM.¹⁰Figure 13.16 F-actin (green) stained by FITC-Phalloidin. Red color is due to autofluorescence of the diatom chloroplasts. Scale bar equals 5 μm.¹¹Figure 3.17 Change of circular structures and space competition with chloroplasts of locomoting diatoms after encountering obstacles. (a-c) a diatom was approaching an obstacle, in which circular structures were clearly seen and chloroplasts was stained with red color. (d-f) Since the diatom can not move the obstacle, it chose to return and the circular structure was squeezing against the chloroplasts to get itself backward.Figure 13.18 Z-stack scanning of a diatom from bottom plane (Z axis coordinate is 1.82 μm) to top plane (Z axis coordinate is 7.60 μm), corresponding from (a) to (d).Figure 13.19 Mucilage trails under SEM, in which pits could be clearly seen in (b) and (c). Scale bars equal to 5 μm (a) and 1 μm (b, c).¹²Figure 13.20 EDAX of mucilage trails and silicon substrate.¹³Figure 13.21 Raman spectra of mucilage trail. SM denotes the spectra from secreted mucilage, while TM-1, TM-2, and TM-3 represent the spectra from three representative mucilage trials.Figure 13.22 Lattice map of adhesion of the mucilage trail.Figure 13.23 Topology of a piece of mucilage trail under AFM Scanning: (a) & (b) are two randomly selected sections.Figure 13.24 Typical AFM measurements in culture medium (a) and mucus (b).Figure 13.25 (a) Force curve against mucilage trails, (b) force curve against EPS.Figure 13.26 Schematic diagram of return process after contact between probe tip and wall. (a) A typical force curve against the trail mucilage, (b) a typical force curve against EPS.Figure 13.27 Schematic diagram of polymer morphology of the mucus (a) and the EPS (b).Figure 13.28 Function of the mucus.Figure 13.29 Scheme of a cycle of repeatable steps (a-e) when a diatom is locomoting to the right in VW model. Briefly, two circular structures alternately use actins as anchors and changing their positions inside the diatom thus generate forces to achieve the locomotion, please refer to the detail description in the text.Figure 13.30 The processes on the cell membrane of diatom contacting the frustule at the raphe.¹⁶Figure 13.31 Locomotion trajectory of several diatoms. Each number represents the trail of a separate diatom.Figure 13.32 Determination of locomotion angle for each step. Positions 1, 2, and 3 represent 3 consecutive positions observed for a motile cell. When diatoms reach position 2, 15 s after position 1, the path direction is defined from the center point of the cell at each time point. After the cell moved to position 3 after another 15 s, the angle of turning from position 2 to position 3 was defined as α, in which α > 0 was defined as turning in a clockwise direction as observed.Figure 13.33 Distribution of locomotion angle.Figure 13.34 Distribution of locomotion length.Figure 13.35 Histogram of locomoting rate of diatoms on different walls and the velocity distributions.Figure 13.36 Comparison of secretion amount of diatom mucus: (a) mucus (b) EPS. Scale bar in (a) 5 μm.¹⁸Figure 13.37 Diffusion and dissolution of diatom secretion.Figure 13.38 Comparison of simulation results (b) and measured results (a) of diatom locomotion trajectory.¹⁹Figure 13.39 Locomotion trajectories of diatoms under different perception parameters: (a) Perception angle = 0°, perception threshold = 1, (b) Perception angle = 0°, perception threshold = 5, (c) Perception angle = 45°, perception threshold = 1, (c) Perception angle = 45°, perception threshold = 5, (e) Perception angle = 90°, perception threshold = 1, (d) Perception angle = 90°, perception threshold = 5.Figure 13.40 Simulated results of diatom adhesion from NetLogo. (a) very strong aggregation, which VMR>20, (b) strong aggregation, which 20>VMR>10, and (c) moderate aggregation, which 10>VMR>1.Figure 13.41 Comparisons between simulated and experimental values of VMR under different perception thresholds either with (a) or without (b) the introduction of cell division, using different perception angles. The two horizontal dashed lines correspond to the 95% confidence intervals from the solid line representing the VMR value determined directly from diatom cultures.Figure 13.42 Simulation results of diatom adhesion under different conditions with proliferation: (a) Perception angle = 0°, perception threshold = 1, (b) Perception angle = 0°, perception threshold = 5, (c) Perception angle = 45°, perception threshold = 1, (d) Perception angle = 45°, perception threshold = 5, (e) Perception angle = 90°, perception threshold = 1, (f) Perception angle = 90°, perception threshold = 5, (g) Perception angle = 135°, perception threshold = 1, (h) Perception angle = 135°, perception threshold = 5.Figure 13.43 The movement trend of daughter cell diatoms after cell division: (a-c) cells stayed in place; (d-f) cell migrated away after cell division.²⁰

14 Chapter 14Figure 14.1 A whimsical view of diatom motility, “Cymbella, Epithemia and Licmophora” by the late Steve Edgar [14.95].Figure 14.2 Diatoms somersault via protruding muscles (1753). The oat-animal, the diatom Craticula cuspidata [14.308], with its two “muscles” protruding from the two ends was described having movements [14.20] analogous to somersaulting [14.108] (reprinted with permission of and designed by Freepik Company; Portrait of naturalist Henry Baker (1698-1774) [14.415] under the Creative Commons Attribution 4.0 International license).Figure 14.3 Andrew Pritchard, naturalist and microscopist [14.408] (1804-1882). 1843 daguerreotype by A.F.J. Claudet [14.351] (public domain image).Figure 14.4 Jean Baptiste Bory de Saint-Vincent, naturalist (1778-1846) [14.406] (public domain image).Figure 14.5 Pierre Jean François Turpin, artist and botanist (1775-1840) [14.404] [14.407] [14.410] (reprinted under the Creative Commons Attribution-Share Alike 4.0 International license).Figure 14.6 The waving fibril model for diatom motility of Robert Jarosch (1962). “Figures 1 to 4 illustrate the relation between the direction of movement of the submicroscopic transverse waves in the hypothetical protoplasmic fibrils and the consequent gliding movements. Figure 1. A single free fibril: the waves running in one direction (W) cause the shifting of the fibril in the opposite direction (P). Figure 2. Longitudinal section through a Chara cell: the undulations (W) of the fibrils, which are fixed to the cell wall (C), cause the shifting of the inner layer of protoplasm (P). Figure 3. Longitudinal section through a gliding organism: (W) direction of waves in extramembranous surface-fixed fibrils; (P) direction of gliding; (M) mucilage; (S) substrate. Figure 4. Direction of waves in extramembranous fibrils during a contraction of Bacillaria paradoxa: (A) an individual cell; (S) substrate” [14.177] (reprinted with permission of Robert Jarosch. Recent photo of Robert Jarosch courtesy of Angelika Jarosch and Ilsa Foissner).Figure 14.7 Diatoms crawl like snails (1838). The motile pennate diatom Epithemia smithii [14.352] as a snail, as described by Christian Gottfried Ehrenberg (1795-1876) in 1838 [14.98]. Right: Christian Gottfried Ehrenberg, naturalist, painted by Eduard Radke ca. 1855 [14.410] (public domain image). Left: His critic, Carl Nägeli, botanist (1817-1891)[14.409] (public domain image).Figure 14.8 The diatom motor is a jet engine (1849). Top: The diatom Gomphonema acuminatum [14.190] [14.348] (scale bar 10 µm) as a jet engine [14.69] (open access; not subject to copyright restriction; the latter reprinted with permission under a GNU Free Documentation License, Version 1.2). Middle left: Aeronautics engineer Francis Herbert Wenham (1824-1908) in 1866 [14.414] (public domain image). Middle right: Lothar Hofmeister, botanist and cell physiologist (1910-1977) [14.195] (reprinted with kind permission of P. Amand Kraml, Director, Observatory of Kremsmünster). Bottom: Diatomist Hamilton Lanphere Smith (1819-1903) [14.350].Figure 14.9 (a) Lesley Ann Edgar, diatomist (1955-2006) [14.65] (image reprinted with permission of Taylor & Francis). (b) Jeremy Pickett-Heaps in 1990 [14.281].Figure 14.10 Jabez Hogg (1817-1899), ophthalmic surgeon [14.260] (reprinted with kind permission of Andrew Tucker, Assistant Curator, Museum of Freemasonry, ©Museum of Freemasonry, London, UK).Figure 14.11 Rowing diatoms (1855). Top: “Quadremes were powerful warships with two banks of oars and multiple rowers per oar” [14.277]. Middle: The diatom oars seen by Jabez Hogg (1855) [14.161]. Next: Mucilage protruding from a raphe of Navicula cuspidata in an SEM of a critical point dried cell [14.90] (reprinted with permission of Springer Nature), scale bars 10 and 1 µm, contrast enhanced by histogram equalization [14.301]. Bottom left: “Navicula confervacea:… Raphe with organic material: valve without marginal spines,” scale bar = 1 µm [14.307] (reprinted with permission of John Wiley and Sons). Bottom right: Scanning electron micrograph showing a single file row of secreted fibrils in purported Pinnularia viridis. Scale bar 0.5 µm. (From [14.154] with permission of John Wiley and Sons). Note that the fibrils are not adjacent to one another, suggesting that they come out of the raphe below individually. See also [14.156].Figure 14.12 Bacteriologist Émile Pierre-Marie van Ermengem (1851-1932) [14.412] (public domain image).Figure 14.13 “Transverse section showing general cellular organization in the region of the pyrenoid [py]. Chloroplast (cp), region of the girdle (gi), intra-pyrenoid lamellae (1), mitochondria (m), nucleus (n), nucleolus (no), raphe (r)” [14.359] (reprinted with permission of Springer Nature). Note a single fibril in the top raphe extending inside past a gap to the cell membrane, below which is a pair of microfilament bundles. The distance between opposite raphes in Cocconeis diminuta is 3 µm [14.359]. Similar gaps have been imaged also by TEM [14.80].Figure 14.14 Left: Max Schultze, microscopic anatomist (1825-1874) [14.405] [14.417] (public domain image). Right: Theodor Wilhelm Engelmann (1843-1909), botanist, physiologist, and microbiologist [14.424] (public domain image).Figure 14.15 Diatoms have protoplasmic tank treads (1865). Top left: The diatom as a double tank tread in girdle view. In order to ensure independence of the two raphes, two tank treads would be necessary. (Adapted from [14.286]. Bottom left: The double tank tread model for diatom motility (1893): “I interpret these phenomena in such a way that a current of cytoplasm is driven through the pole cleft of the anterior end node into the outer cleft of the raphe, there is shifted towards the center and flows back into the cell interior through the outer central node channel” [14.252]. A similar diagram is shown in [14.44]. Right: Georg Ferdinand Otto Müller (1837-1917) [14.419] (public domain image) published a series of 8 papers on diatom motility [14.251] [14.252] [14.253] [14.254] [14.255] [14.256] [14.257] [14.258] [14.387].Figure 14.16 Left: In 2015: “We propose a model [for gliding of Flavobacterium johnsoniae] in which a pinion, connected to a rotary motor, drives a rack (a tread) that moves along a spiral track fixed to the rigid framework of the cell wall. SprB [a cell-surface adhesin], carried by the tread, adsorbs to the substratum and causes the cell to glide…. Tethered cells pinwheel around a fixed axis, suggesting that a rotary motor that generates high torque is a part of the gliding machinery [14.334]…. If 90 nm is the radius of a pinion rotating 3Hz (the maximum speed of rotation when a cell is tethered) then that pinion can drive a rack (a tread carrying adhesins) at 1.5 µm/s, which is the speed that cells glide. This suggests that nature has not only invented the wheel, it also has invented the likes of a microscopic snowmobile…. A model of the gliding machinery. (a) A cross-sectional view of a cell with a rotary gliding motor (blue), a mobile tread (green), a stationary track (red), and an adhesin (magenta). The rotary motor and the track are anchored to the peptidoglycan (PG), and the track is wound spirally around the cell. The rotary motor drives a pinion that engages a mobile tread (rack) that slides along the track. The adhesin, SprB, is attached to the tread and moves with it. The dimension d is the distance between the axis of rotation of the motor and the center of the track, and r is the radius of the pinion. (b) A side view of a cell with a rotary motor powering the motion of a tread carrying SprB” [14.332] [14.333]. O.M. = outer membrane, C.M. = inner cell membrane. Right: A rack and pinion.Figure 14.17 Model for a diatom that moves but leaves no trail [14.75]. Of course, diatoms also lay down the trail.Figure 14.18 Diatoms as the Flame of Life: Capillarity (1883). (a) A candle draws up molten wax by capillarity into the wick, where it burns off along the length of the wick, emitting light as it enters the gaseous phase and ignites. For a diatom the raphe is a hydrophobic wick drawing hydrophobic raphe fluid (raphan) from inside the diatom. As raphan consists of polysaccharide fibers, particles can stick to them as they exit and move along the raphe, carrying the particles along the raphe. If the raphe fluid adheres to a large object or a substrate, the diatom moves in the opposite direction [14.122]. As the raphan comes into contact with water, it hydrates, can no longer wet the raphe, and exits to the medium. The diatom trail is analogous to the flame of a candle. (b) “The capillarity model for diatom gliding locomotion, as originally conceived in [14.122]. A schematic ‘longitudinal slice’ of a single raphe is shown as if the raphe went straight through the silica valve. (It is actually hooked in cross section.) The crystalloid bodies empty their fibrillar, mucopolysaccharide contents into the raphe via exocytosis. The role of the microfilaments was speculated to be control of the distribution of the exocytosis along the raphe in an unspecified manner. The directionality of the motion could come either from an asymmetric distribution of release of mucopolysaccharide along the raphe, or, as shown here, from a postulated difference in rate of hydration of the mucopolysaccharide between the leading and trailing pores. The hydration of the mucopolysaccharide may also permit it to stick to the substratum, as indicated. This results in motion in the direction shown. The released, fully hydrated mucopolysaccharide stays attached to the substratum as trail material. While capillarity fills the raphe with the mucopolysaccharide, its hydration removes it and provides the driving force. (Hydrated mucopolysaccharide no longer wets the walls of the raphe, suggested to consist of a hydrophobic lipid layer by [14.92]” [14.117] (reprinted with permission of Elsevier). The sketch has been rotated and aligned to correspond to the candle. If the substrate were replaced by small particles, they would rise upwards just as the liquid wax does in a candle. The sketch has been modified to show how the microfilament bundles may permit access of the crystalloid bodies [14.79] at one end of the raphe but not the other, by sliding along their length. Note: “We have electron micrographic evidence that indeed these vesicles are secreted into the raphe canal” [14.47]. In this model, the motile pennate diatom is the flame of life.Figure 14.19 Bellowing diatoms (1887). Left: Stauroneis baileyi, shown in girdle view, was found to bellow out, with the distance between a and b increasing, when moving in the direction of the arrow [14.342]. Right: A selfcompressing bellows, on blowing air out, would move in the same direction [14.106] (public domain image).Figure 14.20 Top: In 1896: Cross sections of the pennate diatom Pinnularia major from the middle to the end of the cell showing how the raphe is a slit through the whole valve [14.205]. Bottom: Robert Lauterborn in 1928 [14.239] (reprinted with permission of Elsevier).Figure 14.21 Jelly powered jet skiing diatoms (1896). In 1892: “… Pinnularia nobilis in motion in the view on the girdle side. n the nucleus, c the centrosome, x the peculiar double threads in the plasma, a the inflow to the node (k) of the anterior raphe, b the gelatin thread that shoots out to the rear, which has rolled up at the right end at the end. The arrows indicate the direction of movement of the diatom, the inflow and the thread” (translated from [14.44]). Otto Bütschli (1848-1920) [14.418] attributed the diatom’s motion to pulling, pushing and recoil of the threads, rather than the inward flow he thought he observed. (Photograph of Bütschli by Max Kögel [14.191] with permission of Universitätsbibliothek Heidelberg under the Creative Commons Attribution-Share Alike 4.0 International license).Figure 14.22 Bubble powered diatoms (1905). Left: The bubbly motile diatom. Middle: Nitzschia acicularis [14.368]. Right: View of a portion of the Chemical Laboratory at the Mt. Prospect Laboratory of the Brooklyn Water Works, next door to the Biological Laboratory, where Daniel D. Jackson [14.80] worked [14.403].Figure 14.23 Diatoms win: “I have no new theory to offer and see no reason to use those already abandoned” (1940) [14.230]. Left: Medical botanist Pierre Martens (1895-1981) in 1950 [14.73]. Right: Martens’ depiction of Otto Müller’s flow model: “Connective view of a valve, showing the channeling at the level of the median nodule and the paths attributed to the propellant currents. The median nodule is drawn (after Müller), in an orientation which does not show the outlet of the ‘external slit’ outside; the internal structure of the polar nodules is not shown. The upper arrow indicates the direction of locomotion…. Abbreviations: np = polar nodule; nc = central nodule; fe = external slot of the raphe; fi = internal slot; rc = groove of the central nodule, joining the two internal slots of the same valve; cu = small channel joining the external and internal slits”¹³ [14.230].Figure 14.24 Is diatom motility a special case of cytoplasmic streaming? (1943). Left: Early model of what we now call a motor protein invoked for cytoplasmic streaming. 1) Protein molecule in extended form. Bond at anterior end of molecule. (2) Protein molecule in contracted form. (3) Shifting of bond from anterior to posterior end. (4) Protein molecule in expanded form…. Let us say that the molecules which form bonds in this way so as to contribute to the motion are in phase and those which may form bonds so as to oppose the motion are out of phase. It is clear that streaming can occur only when the number of molecules in phase exceeds those out of phase. This cannot happen by chance alone and we must therefore postulate that some mechanism is present which sets the majority of stream proteins in phase. This mechanism must of course be intimately related to the mechanism responsible for the reversal of streaming” [14.222] (reprinted with permission of the American Philosophical Society). Right: Motion of an inchworm caterpillar [14.287] (reprinted with permission of Elsevier).Figure 14.25 “Row, row, row your boat. The molecular motors in the myosin heads pull the myosin filaments (blue) over the actin filaments” [14.157]. This version from [14.46] (reprinted with permission of Cambridge University Press). Cf. Figure 14.11.Figure 14.26 Model for an amoeba (1999) [14.10] (reprinted with permission of Elsevier).Figure 14.27 In 2004: “A schematic representation of a likely mechanism underlying the fluctuation enhancement of an actin filament, induced by Chara myosin molecules. (a) Disequilibrating state: A myosin head generating the sliding force influences the kinetics of the neighboring heads along the actin filament, in which either the pushing or the pulling force is generated. (b) Coordinating state: These activated heads develop a coordination among themselves along the filament. Coordination in one place then induces a disequilibrium out of coordination in the neighborhood. Equilibration and desequilibration thus reiterate” [14.144] (reprinted with permission of Elsevier).Figure 14.28 “Schematic picture of the delivery of different cell wall components to the plant plasma membrane…. Question marks indicate that no direct experimental evidence for the localisation of the component is available…. The vesicles containing the CESAs [cellulose synthase proteins] are transported to the plasma membrane with the help of actin cables” [14.111] (reprinted with permission of Elsevier). A myosin XI such as in the class of those involved in cytoplasmic streaming functions in cellulose exocytosis [14.441].Figure 14.29 Top left: “L.S. [lateral section] in valve view (Navicula cuspidata) showing detail of microfilamentous bundles and associated vesicles; some vesicles appear attached to the bundles (arrow). Scale = 1 µm” [14.93] (reprinted with permission of Elsevier). These are presumed to carry raphe fibrils (raphan). Top right: An F-actin = filamentous actin = microfilament can carry secretory vesicles to the cell membrane [14.337]. In this depiction, transport of vesicles can be either by direct attachment to myosin or indirectly via hydrodynamic flow [14.42] (reprinted with permission of John Wiley & Sons). The attachment of myosin to a vesicle may involve “membrane-anchored core myosin receptors, possibly aided by adaptors” [14.279]. Myosin XI/vesicle attachments are transient, lasting only a few seconds [14.30]. Middle left: An explicit model for binding of myosin to a membrane, in this case the outer (ONM) and inner (INM) nuclear membranes [14.442] (reprinted with kind permission of Iris Meier). Bottom: “The microfilament model for diatom gliding locomotion, as conceived by Edgar & Pickett-Heaps (1984) [14.93], and adapted from their Figures 39, 40 and 41. All features are the same as in [Figure 14.18], except that: (a) the mucopolysaccharide fibrils are assumed to be attached to the microfilament bundles through the plasmalemma, which has no effect because of its fluid nature, via “the lining of the vesicles [crystalloid bodies] in which the strands were originally synthesized”; (b) hydration is assumed to occur along the whole length of each mucopolysaccharide fibril while it is still within the raphe; (c) the fibrils are assumed to swell and elongate as they come out of the crystalloid bodies; (d) a mechanism is needed by which “the mucilage strands are broken free from the plasmalemma on reaching the apical raphe ending [trailing pore].” The microfilament bundle is presumed to provide the motive force. Reversal of the direction of motion is suggested to occur as follows: “Since... there are two bundles of filaments..., perhaps the actin filaments in each bundle are oriented in one direction, and the polarities in the two bundles of filaments are opposite. If this were so, bidirectional movement could be envisaged as occurring as...the raphe adhesive is moved by some controlled activation of or coupling with the actin bundles alternately.” Only one microfilament bundle is shown. The other, oppositely oriented, would be placed behind this one. Thus, two rows of fibrils are possibly present in a raphe simultaneously. The second row would be behind the one shown” [14.117] (reprinted with permission of Elsevier).Figure 14.30 Diatom adhesion as a sliding toilet plunger (1966). Left: “Suction cup with handle, used to study suction-seal-substratum relationships” [14.79] (reprinted with permission of Springer Nature). Right: Ryan W. Drum in 2003.Figure 14.31 Diatom as a monorail that lays its own track (1967). Top: In 1967: “Either of the mechanisms indicated…could keep the trail against the raphe and explain adhesion forces. In the…osmotic… hypothesis…(a), the trail T is held in position by low pressure in the water W in the raphe slit. The pressure would be kept low by osmosis across the cell membrane. Alternatively,…the interfacial tension hypothesis… (b) shows the raphe filled with a liquid L immiscible with water. Interfacial tension at I would then keep the trail in place…. Schematic raphe cross-sections. Substrate stippled. I, water-liquid interface; L, liquid; T, trail; W, water at low pressure being sucked into the diatom by osmosis” [14.142] (reprinted with permission of Taylor & Francis). Middle right: A monorail train gripping its track [14.309] (public domain image). If diatoms do indeed lay down their own track, we have much to learn from them [14.430]. Bottom left: Margaret A. Harper and John F. Harper, wedding photo taken at Willesden Parish Church, NW London, UK, 29 August 1964. Bottom right: Margaret A. Harper and John F. Harper, at present, with their permission.Figure 14.32 “Four locomotion theories [as of 1977], represented on schematic sections of pennate diatoms in their apical planes. Trail precursor is shown by coarse stipple, trail by fine stipple. (a) Jarosch (1962) [14.177]. Mucilage secreted at pores a and c being driven by undulating actin filaments connected to the protoplast. (b) Hopkins & Drum (1966) [14.164]. Trail secreted through entire lengths of raphes ab and cd and expanding on leaving the raphe system. (c) Harper & Harper (1967) [14.142]. Motion due to trail secretion from pore a. Secretion from upper pores e and h forming lump at l and particle p being carried forward. (d) Gordon & Drum (1970) [14.122]. Capillary flow of a liquid along raphes ab and cd, conversion to trail at pores b and d” [14.141] (reprinted with permission of John Wiley and Sons). Note that the central nodule is lacking in diatoms such as Bacillaria [14.439], which either reduces the complexity from four pathways, as in Liebsch (1929) [14.218], or indicates that the central nodule does not play the role suggested here.Figure 14.33 The diatom as a “compressed air” Coanda Effect gliding vehicle (1967). A wing profile assisted by compressed air coming out of the plenum chambers, with the air flowing close to the surface due to the Coanda effect [14.22] (reprinted with kind permission of Stephen D. Prior, superimposed by diatom Cymbella cistula [14.244] with permission of the Muséum national d’Histoire naturelle under a Creative Commons license). Of course, the raphes should be rotated 90° for a proper match.Figure 14.34 The electrokinetic diatom (1974). Depiction of ionic currents that break the symmetry of the spherical Fucus egg [14.172] (reprinted with permission of Elsevier). Such currents, if they occur in pennate diatoms, could bias the flow of charged molecules in the raphe or the cell membrane just within the raphe or the microfilaments adjacent to the raphe. Reversal of the field would then be predicted when the diatom reverses direction.Figure 14.35 Discovery of the “crystalloid bodies” and their relationship to the raphe. Left: “1. A three-dimensional view of the silica structure of a diatom. The raphe system (RS) is composed of a central pore (CP) and a terminal pore (TP) with an outer groove of the raphe fissure (R). Continuation grooves (CG) and a probable anterior pore (AP) are shown. 2. A raphe plane section of a diatom moving upon a plane surface (GS) showing the four pores as in #1. The cytoplasm contains a fibrillar bundle (FB) and crystalloid bodies (C) containing minute fibrils (F). These are found in the diatom raphe system and in the diatom trail. The point of locomotor-adhesion contact (LAC) is indicated. 3. A raphe plane section of a diatom moving upon particles (PS). In this case, locomotor-adhesion contact (LAC) can involve any point along the raphe system” [14.164] (reprinted with permission of Taylor & Francis). Right: “Transverse cross-section of a raphe fissure and adjacent cytoplasm of a diatom moving over a particulate substratum; the locomotor adhesion seal (arrow), firmly attached to particles, is being pushed along raphe by streaming directed against that seal; the fibrillar bundle (F) lies next to the raphe (R); longitudinal and transverse sections of crystalloid bodies (CB) are also shown” [14.79] (reprinted with permission of Springer Nature). The fibrillar bundles were later identified as similar to smooth muscle and presumed to contract, pushing the crystalloid bodies and moving the diatom [14.173].Figure 14.36 The diatom clothes line or railroad track (1980). Left: Clothespin line model for diatom motility, amalgamated from [14.11] [14.427] (public domain image). Top to bottom: microfilament bundle, myosins, clothespins are membrane bound raphan synthase carrying raphan inside a raphe, where raphan is a polysaccharide raphe fibril. Top right: “Model for the organization of the motor apparatus in diatoms. Adhesive mucilage secreted by the diatom adheres to the substrate and binds to as yet undefined transmembrane components. The cytoplasmic domain of the membrane-associated complex is linked to a diatom myosin which actively translocates the membrane complex and attached mucilage rearward along a track of cortical actin filaments, leading to forward gliding of the diatom” [14.146] (reprinted with permission of Elsevier). This is the generally assumed model of Lesley Edgar and Jeremy Pickett-Heaps, though they also allowed for an indirect coupling of the myosin to the raphe fibrils [14.93]. Bottom Right: Similar model of Rick Wetherbee et al. (1998): “Diagram suggesting how the adhesion complex would look in a raphid diatom. Note that only the actin [microfilament] has been described for certain. There has been no attempt to illustrate where components responsible for motility (e.g., a motor) might be located” [14.401] (reprinted with permission of John Wiley & Sons). The actin-associated proteins would presumably be myosins, and the transmembrane proteins would be raphan synthase.Figure 14.37 Diatom ion cyclotron resonance (1987). A take on diatoms and cyclotrons [14.207] (public domain image). Accelerating diatoms are Cymbella cistula [14.244] (reprinted with permission of the Muséum national d’Histoire naturelle under a Creative Commons license).Figure 14.38 Diatoms do internal treadmilling (1998). Left: In treadmilling of microfilaments, actin monomers are added at one end and removed at the other end, resulting in a constant length, with motion [14.45] (reprinted with permission of Springer Nature). Upper right: Molecular motors may be involved in regulating the length of a treadmilling microfilament: “A treadmilling filament is described by a lattice of dynamic length. Motors are represented as particles that occupy the sites. At one end sites are added at a rate of α. At the opposing end, empty (occupied) sites are removed at a rate of β . Particles attach to empty sites at a rate of ω and detach from the lattice at a rate of . Particles hop to adjacent free sites in the direction of the shrinking end at a rate of γ” [14.100] (reprinted with permission of the American Physical Society Publishing). Lower right: “A model showing how myosin driven actin sliding with the combination of tethering proteins can potentially drive ER [endoplasmic reticulum] and Golgi mobility. Myosins are shown linking actin filaments within a bundle and are responsible for filament sliding” [14.234] (reprinted with kind permission of Joseph F. McKenna). Similar sliding might be occurring in the microfilament bundles adjacent to raphes.Figure 14.39 Flow along a line, such as a raphe, induces a circulating flow of the adjacent fluid, by analogy to the early neural plate in a chicken [14.402] (reprinted with permission of Springer Nature).Figure 14.40 Rough schematic for a diatom robot (diatombot), a neutrally buoyant, remotely controlled robot that simulates motion of the raphe fluid using bungee cords moving on pairs of motorized pulleys to test the possibility that motile diatoms can swim. The outside portions of the bungee cords lie in grooves, allowing them to make contact with the water outside. Particles in the water could show up in any induced water flows.Figure 14.41 Surface treadmilling, swimming and snorkeling diatoms (2007). A snorkeling pennate diatom.Figure 14.42 Acoustic streaming: the diatom as vibrator or jack hammer (2010). Perhaps a vibrating diatom can smash its way through sand, like a jack hammer removes concrete [14.306] (reprinted with kind permission of Robert Bosch Tool Corporation). The diatom is Lyrella esul [14.338] (reprinted with kind permission of David A. Siqueiros Beltrones).Figure 14.43 Propulsion of diatoms via many small explosions (2020). Nuclear propulsion via hundreds of “small” nuclear bombs: Project Orion (conceived by Stanislaw Ulam, 1946), an analog of diatom propulsion via explosive hydration of raphe fibrils. Top left: artist’s conception [14.420]. Top right: Principle [14.315]: The “Pulse unit injection” would correspond to a raphe carrying raphe fibrils that explosively hydrate on being exposed to water. Middle right: A space faring diatom [14.119] [14.124] Lyrella esul propelled by explosions [14.134]; Lyrella esul [14.338] with kind permission of David A. Siqueiros Beltrones. Bottom: Project Orion configuration [14.420], all others from NASA, in public domain).Figure 14.44 Diatoms walk like geckos (2019). Was the gecko inspired by the diatom, or vice versa? (Gecko [14.206] . Pseudoraphid diatom Podocystis adriatica [14.139] with permission of Paul Hargraves and the New England Botanical Club).Figure 14.45 Intraprotein pores in hyaluronan synthase and cellulose synthase. Top left: “A model for interaction of the HAS [hyaluronan synthase] tandem-motif region with HA [hyaluronan, the #s refer to membrane domains]. The… growing HA-UDP chain, containing alternating GlcNAcβ1,4 (blue squares) and GlcUAβ1,3 (blue-white diamonds) attached to UDP [uridine diphosphate] (red inverted triangle) at the reducing end. Preliminary results indicate that the nonreducing end contains a chitin oligomer cap with four GlcNAc residues, which is the primer on which HA synthesis is initiated” [14.18] (reprinted with permission of Oxford University Press). Top right: AFM experiment suggesting that raphe fibrils are multistranded [14.153] (like cellulose) (reprinted with permission of Elsevier). Bottom: “Updated models of plant CESAs [cellulose synthases] and CSCs [cellulose synthesis complexes]. (a) A computational model of a plant CESA catalytic domain with P-CR and CSR regions (light grey). The glucan chain (purple) is from the homologous Rhodobacter structure. The location of the transmembrane helices (TMH) is represented with grey boxes. (b) Three CESAs, encoded by three different genes, may interact to form a trimeric particle, which in turn may assemble into a hexameric rosette, depicted in (c). The glucan chains are represented in red. (Image (a) is adapted from [14.339] and is courtesy of Jonathan Davis)…. the intra-protein tunnel [pore]… provides a low-energy pathway for translocating the growing glucan chain to the external membrane surface from the cytoplasmic side, where the catalytic site transfers a glucose residue from UDP-glucose to the reducing end of the glucan” [14.62] (reprinted with permission of Elsevier).Figure 14.46 Left: “Stages in secretion at the raphe (hypothetical). (a) Polysaccharide fibrils are discharged from a vesicle into the raphe at the central pore. The preferred secretory sites are at both central and apical (not shown) raphe endings; (b) Secreted fibrils experience hydration and begin to swell and elongate. A second vesicle is docked at the plasmalemma and primed for exocytosis. (c) After considerable swelling the mucilaginous strands project through the raphe; their proximal ends are attached to the plasmalemma and their distal ends are free to make contact with a substratum; (d) Strands, which were produced at the raphe ending, have moved along the raphe but still occupy the slit and maintain attachment to the plasmalemma. Microfilaments act as a barrier preventing vesicle discharge here (not evident in all cells)” [14.93] (reprinted with permission of Elsevier). Note that the hydration of the raphe fibrils does not likely occur within the raphe, as shown, because the walls of the raphe are hydrophobic [14.92] [14.93] [14.122]. On the contrary, hydration at the external tip would generate a force pulling the fibrils out of the raphe. Right: The silica lining the raphe differs from that of the valve [14.66] (reprinted with permission of John Wiley & Sons).Figure 14.47 A general model for noncellulosic polysaccharide biosynthesis [14.362] (reprinted with permission of Oxford University Press).Figure 14.48 “TEM of thin sections of Amphora veneta. 26: T. S. [thin section] through a perinuclear dictyosome [Golgi]. The dictyosome is composed of 8 cisternae (ct) which are blebbing fibrous (v₁) and smaller granular (v₂) vesicles (× 44,000). 27: Oblique L. S. through the central region. Irregularly shaped fibrous (v₁) and smaller granular vesicles (v₂) are apparent (× 32,400). 28: T. S. part of cell to show frustule-cell-membrane interface (fcm). Large numbers of vesicles (v₄) are present outside the cell membrane (cm) but within the silica wall (sw) (× 33,000). 29: T. S. showing fibrous material (fm) extruding from a raphe fissure (rf) (× 22,400)” [14.70] (reprinted with permission of Springer Nature). Note: “The largest [vesicles] (27), were derived from the mature dictyosome [Golgi] cisternae, and were often irregular in outline whilst containing fibrillar material. These vesicles were widely distributed throughout the cytoplasm and the contents were similar in morphology to the extracellular mucilaginous material (29).”Figure 14.49 “Four possible configurations of the microfilament bundles, assuming that they are capable of sliding a short distance relative to the silica raphe. Only the ends of the raphe are shown here, represented by two pores. The view is from the inside of the cell, looking out. The placement of the raphe pores on opposite sides of the apical axis [14.34] is a common feature of raphid diatoms. In this scheme, the microfilament bundles block access of the crystalloid bodies to the raphe along their whole length. In the capillarity model, such blockage would explain the direction and reversal of motion, as indicated on the right” [14.117]. (Reprinted with permission of Elsevier).Figure 14.50 Top: Swirling, draining patterns on a bubble film draining downwards [14.250] (reprinted with kind permission of Michael Reese Much FRMS). Bottom: One frame from a movie showing how dynamic these patterns are [14.288]. The interference colors are due to varying thickness. Note that the flows organize themselves along narrow lines which can draw in material from a wide area. This could be analogous to a diatom’s raphe drawing the liquid cell membrane towards it, bearing proteins (raphan synthase) ready to secrete their polysaccharide raphe fibrils (raphan) upon reaching the raphe.Figure 14.51 Surfing diatoms, Achnanthes and Podocystis [14.95].Figure 14.52 Membrane surfing: A new working hypothesis for the diatom motor (2020). Surfing working model for diatom motility. Raphan synthase is a hypothesized membrane protein that is functional either in the cell membrane, or more likely, in vesicles (crystalloid bodies) into which it deposits raphan, the presumed polysaccharide constituting the raphe fibrils seen in raphes. The whole cell membrane flows, bringing the raphan synthases or the crystalloid bodies, represented as miniatures of Figure 14.51, to the raphe. Myosin motor molecules move along the microfilament bundles (only one shown) hydrodynamically inducing flow of the cell membrane by carrying the vesicles over the raphe. As the vesicles fuse with the cell membrane, they dump their hydrophobic contents into the raphe, where the hydrophobic raphan is shown as short, vertical lines. As the raphe lining is hydrophobic, they fill the raphe via capillarity. Those that come in contact with a substrate, such as the chondrite [14.125] shown here, hydrate, swell, and stick to it. As these hydrated raphan molecules exit, more hydrophobic raphan molecules fill in the raphe, causing a net flow of the anhydrous raphan within the raphe, shown by the small arrow. The result is that the diatom moves relative to the chondrite in the opposite direction, shown by the large arrow. On a flat substrate, the swollen raphan fibrils are left behind as the diatom trail. The flow of the cell membrane determines the direction in which the diatom moves relative to its substrate. Capillarity provides the tremendous force. “On Earth, capillary forces have to fight gravity. But in space, the only resistance is the viscosity of the liquid, which slows the flow but cannot stop it” [14.240]. At the size scale of the diatom raphe, gravity has negligible effect, so the slogan applies.Figure A14.1 Could squeezing out of raphe fluid cause diatom motility? Photo of caulking gun [A.14.4] superimposed with a Cymatopleura diatom [A.14.1] under the Creative Commons Attribution License (CC BY 4.0).