Diversity Takes Shape: Understanding the Mechanistic and Adaptive Basis of Bacterial Morphology

<div><p>The modern age of metagenomics has delivered unprecedented volumes of data describing the genetic and metabolic diversity of bacterial communities, but it has failed to provide information about coincident cellular morphologies. Much like metabolic and biosynthetic capabilities, morphology comprises a critical component of bacterial fitness, molded by natural selection into the many elaborate shapes observed across the bacterial domain. In this essay, we discuss the diversity of bacterial morphology and its implications for understanding both the mechanistic and the adaptive basis of morphogenesis. We consider how best to leverage genomic data and recent experimental developments in order to advance our understanding of bacterial shape and its functional importance.</p></div

The Caulobacterales lineage exhibits diversification of the prosthecate morphology.

<p><b>(A)</b> Phylogeny of the order Caulobacterales generated as described in <a href="http://www.plosbiology.org/article/info:doi/10.1371/journal.pbio.1002565#pbio.1002565.g001" target="_blank">Fig 1</a>. Schematics and corresponding colors indicate inferred ancestral morphologies and their subsequent inheritance. Black branches indicate rod-shape, nonappendaged morphology, including several apparent prostheca loss events. Scale bar indicates 0.1 amino acid substitutions per site. <b>(B)</b> Transmission electron micrographs of members of the Caulobacterales, highlighting disparate prosthecate morphologies. For each morphology, a brief description and the name of one representative species is provided, followed by the image source in parentheses. <b>1. Bilateral prosthecae, <i>Asticcacaulis biprosthecum</i></b> (Chao Jiang, Stanford University). <b>2. Subpolar prostheca, <i>Asticcacaulis excentricus</i></b> (Chao Jiang, Stanford University). <b>3. Polar prostheca, <i>Caulobacter crescentus</i></b> (Paul Caccamo, Indiana University). <b>4. Polar prostheca, <i>Maricaulis maris</i></b> (Patrick Viollier, University of Geneva). <b>5. Short polar prostheca, <i>Brevundimonas subvibriodes</i></b> (Brynn Heckel, Indiana University); note other members of this genus display a much longer prostheca. <b>6. Polar prostheca through which budding reproduction occurs, <i>Hirschia baltica</i></b> (Paul Caccamo, Indiana University). Magnification varies between micrographs. All images are reproduced with permission.</p

Myriad morphologies have evolved throughout the bacterial domain.

<p>Bacterial phylogeny derived from genome sequence data for selected species, with an emphasis on morphologically and phylogenetically diverse taxa. Sequence data gathered from the Joint Genome Institute [<a href="http://www.plosbiology.org/article/info:doi/10.1371/journal.pbio.1002565#pbio.1002565.ref003" target="_blank">3</a>] and the National Center for Biotechnology Information [<a href="http://www.plosbiology.org/article/info:doi/10.1371/journal.pbio.1002565#pbio.1002565.ref004" target="_blank">4</a>] were searched for reference genes and aligned using Phylosift [<a href="http://www.plosbiology.org/article/info:doi/10.1371/journal.pbio.1002565#pbio.1002565.ref005" target="_blank">5</a>]. FastTree [<a href="http://www.plosbiology.org/article/info:doi/10.1371/journal.pbio.1002565#pbio.1002565.ref006" target="_blank">6</a>] generated an approximate maximum likelihood tree from the resulting concatenated alignment. The final tree was formatted using iTol [<a href="http://www.plosbiology.org/article/info:doi/10.1371/journal.pbio.1002565#pbio.1002565.ref007" target="_blank">7</a>]. Black dots denote ancestral nodes of selected major taxa: <b>DT</b>, Deinococcus-Thermus; <b>Ac</b>, Actinobacteria; <b>Cf</b>, Chloroflexi; <b>Cn</b>, Cyanobacteria; <b>Fi</b>, Firmicutes (inclusive of Mollicutes); <b>Sp</b>, Spirochetes; <b>PVC</b>, Planctomycetes, Verrucomicrobia, Chlamydiae; <b>Cb</b>, Chlorobi; <b>Bd</b>, Bacteroidetes; <b>α</b>, <b>β</b>, <b>γ</b>, <b>δ</b>, <b>ε</b>, Proteobacteria subdivisions. <b>1. <i>Bifidobacterium longum</i>. 2. <i>Streptomyces coelicolor</i></b> (mycelial [multicellular] filament with hyphae and spores). <b>3. <i>Corynebacterium diphtheria</i>e</b> (two cells, dumbbell and club shapes). <b>4. <i>Herpetosiphon aurantiacus</i></b> (filament of multiple cylindrical cells). <b>5. <i>Calothrix</i></b> (filament of multiple disk-shaped cells). <b>6. <i>Mycoplasma genitalium</i>. 7. S<i>piroplasma culicicola</i>. 8. <i>Lactococcus lactis</i></b> (predivisional cell). <b>9. <i>Borrelia burgdorferi</i></b>. <b>10. <i>Gimesia maris</i></b> (previously <i>Planctomyces maris</i>, predivisional cell with proteinaceous stalk). <b>11. <i>Prosthecochloris aestuarii</i></b>. <b>12. <i>Pelodictyon phaeoclathratiforme</i></b> (filament of multiple trapezoidal cells). <b>13. <i>Spirosoma linguale</i></b>. <b>14. <i>Muricauda ruestringensis</i></b> (appendage includes nonreproductive bulb). <b>15. <i>Desulfovibrio vulgaris</i></b> (two cells, helical and curved shapes). <b>16. <i>Helicobacter pylori</i></b>. <b>17. <i>Caulobacter crescentus</i></b> (predivisional cell). <b>18. <i>Hyphomonas neptunium</i></b> (predivisional cell)<b>. 19. <i>Rhodomicrobium vannielii</i></b> (filament of multiple ovoid cells, one is predivisional)<b>. 20. <i>Prosthecomicrobium hirschii</i></b>. <b>21. <i>Simonsiella muelleri</i></b> (filament of multiple curved cells). <b>22. <i>Nevskia ramosa</i></b> (two cells with bifurcating slime stalk)<b>. 23. <i>Beggiatoa leptomitiformis</i></b> (filament of multiple, giant cylindrical cells). <b>24. <i>Thiomargarita nelsonii</i></b> (single, giant cell). <b>25. <i>Escherichia coli</i>. 26. <i>Mariprofundus ferrooxydans</i></b> (single cell with metal-encrusted stalk). Bacterial schematics are not to scale. Species names are colored according to morphology as indicated in the key. Colored dots are appended to indicate species with multiple morphologies. Names of species depicted in schematics are emphasized in large, bold font.</p

Timescales and Frequencies of Reversible and Irreversible Adhesion Events of Single Bacterial Cells

In the environment, most bacteria form surface-attached cell communities called biofilms. The attachment of single cells to surfaces involves an initial reversible stage typically mediated by surface structures such as flagella and pili, followed by a permanent adhesion stage usually mediated by polysaccharide adhesives. Here, we determine the absolute and relative timescales and frequencies of reversible and irreversible adhesion of single cells of the bacterium <i>Caulobacter crescentus</i> to a glass surface in a microfluidic device. We used fluorescence microscopy of <i>C. crescentus</i> expressing green fluorescent protein to track the swimming behavior of individual cells prior to adhesion, monitor the cell at the surface, and determine whether the cell reversibly or irreversibly adhered to the surface. A fluorescently labeled lectin that binds specifically to polar polysaccharides, termed holdfast, discriminated irreversible adhesion events from reversible adhesion events where no holdfast formed. In wild-type cells, the holdfast production time for irreversible adhesion events initiated by surface contact (23 s) was 30-times faster than the holdfast production time that occurs through developmental regulation (13 min). Irreversible adhesion events in wild-type cells (3.3 events/min) are 15-times more frequent than in pilus-minus mutant cells (0.2 events/min), indicating the pili are critical structures in the transition from reversible to irreversible surface-stimulated adhesion. In reversible adhesion events, the dwell time of cells at the surface before departing was the same for wild-type cells (12 s) and pilus-minus mutant cells (13 s), suggesting the pili do not play a significant role in reversible adhesion. Moreover, reversible adhesion events in wild-type cells (6.8 events/min) occur twice as frequently as irreversible adhesion events (3.3 events/min), demonstrating that most cells contact the surface multiple times before transitioning from reversible to irreversible adhesion

Microfluidic Device for Automated Synchronization of Bacterial Cells

We report the development of an automated microfluidic “baby machine” to synchronize the bacterium <i>Caulobacter crescentus</i> on-chip and to move the synchronized populations downstream for analysis. The microfluidic device is fabricated from three layers of poly­(dimethylsiloxane) and has integrated pumps and valves to control the movement of cells and media. This synchronization method decreases incubation time and media consumption and improves synchrony quality compared to the conventional plate-release technique. Synchronized populations are collected from the device at intervals as short as 10 min and at any time over four days. Flow cytometry and fluorescence cell tracking are used to determine synchrony quality, and cell populations synchronized in minimal growth medium with 0.2% glucose (M2G) and peptone yeast extract (PYE) medium contain >70% and >80% swarmer cells, respectively. Our on-chip method overcomes limitations with conventional physical separation methods that consume large volumes of media, require manual manipulations, have lengthy incubation times, are limited to one collection, and lack precise temporal control of collection times

Physiochemical Properties of <i>Caulobacter crescentus</i> Holdfast: A Localized Bacterial Adhesive

To colonize surfaces, the bacterium <i>Caulobacter crescentus</i> employs a polar polysaccharide, the holdfast, located at the end of a thin, long stalk protruding from the cell body. Unlike many other bacteria which adhere through an extended extracellular polymeric network, the holdfast footprint area is tens of thousands times smaller than that of the total bacterium cross-sectional surface, making for some very demanding adhesion requirements. At present, the mechanism of holdfast adhesion remains poorly understood. We explore it here along three lines of investigation: (a) the impact of environmental conditions on holdfast binding affinity, (b) adhesion kinetics by dynamic force spectroscopy, and (c) kinetic modeling of the attachment process to interpret the observed time-dependence of the adhesion force at short and long time scales. A picture emerged in which discrete molecular units called adhesins are responsible for initial holdfast adhesion, by acting in a cooperative manner

Microfluidic Device for Automated Synchronization of Bacterial Cells

We report the development of an automated microfluidic “baby machine” to synchronize the bacterium <i>Caulobacter crescentus</i> on-chip and to move the synchronized populations downstream for analysis. The microfluidic device is fabricated from three layers of poly­(dimethylsiloxane) and has integrated pumps and valves to control the movement of cells and media. This synchronization method decreases incubation time and media consumption and improves synchrony quality compared to the conventional plate-release technique. Synchronized populations are collected from the device at intervals as short as 10 min and at any time over four days. Flow cytometry and fluorescence cell tracking are used to determine synchrony quality, and cell populations synchronized in minimal growth medium with 0.2% glucose (M2G) and peptone yeast extract (PYE) medium contain >70% and >80% swarmer cells, respectively. Our on-chip method overcomes limitations with conventional physical separation methods that consume large volumes of media, require manual manipulations, have lengthy incubation times, are limited to one collection, and lack precise temporal control of collection times

Physiochemical Properties of <i>Caulobacter crescentus</i> Holdfast: A Localized Bacterial Adhesive

To colonize surfaces, the bacterium <i>Caulobacter crescentus</i> employs a polar polysaccharide, the holdfast, located at the end of a thin, long stalk protruding from the cell body. Unlike many other bacteria which adhere through an extended extracellular polymeric network, the holdfast footprint area is tens of thousands times smaller than that of the total bacterium cross-sectional surface, making for some very demanding adhesion requirements. At present, the mechanism of holdfast adhesion remains poorly understood. We explore it here along three lines of investigation: (a) the impact of environmental conditions on holdfast binding affinity, (b) adhesion kinetics by dynamic force spectroscopy, and (c) kinetic modeling of the attachment process to interpret the observed time-dependence of the adhesion force at short and long time scales. A picture emerged in which discrete molecular units called adhesins are responsible for initial holdfast adhesion, by acting in a cooperative manner

Programmable, Pneumatically Actuated Microfluidic Device with an Integrated Nanochannel Array To Track Development of Individual Bacteria

We describe a microfluidic device with an integrated nanochannel array to trap individual bacteria and monitor growth and reproduction of lineages over multiple generations. Our poly­(dimethylsiloxane) device comprises a pneumatically actuated nanochannel array that includes 1280 channels with widths from 600 to 1000 nm to actively trap diverse bacteria. Integrated pumps and valves perform on-chip fluid and cell manipulations that provide dynamic control of cell loading and nutrient flow, permitting chemostatic growth for extended periods of time (typically 12 to 20 h). Nanochannels confine bacterial growth to a single dimension, facilitating high-resolution, time-lapse imaging and tracking of individual cells. We use the device to monitor the growth of single bacterial cells that undergo symmetric (<i>Bacillus subtilis</i>) and asymmetric (<i>Caulobacter crescentus</i>) division and reconstruct their lineages to correlate growth measurements through time and among related cells. Furthermore, we monitor the motility state of single <i>B. subtilis</i> cells across multiple generations by the expression of a fluorescent reporter protein and observe that the state of the epigenetic switch is correlated over five generations. Our device allows imaging of cellular lineages with high spatiotemporal resolution to facilitate the analysis of biological processes spanning multiple generations

Relevant steps of PG biosynthesis and D-amino acid dipeptide (DAAD) probes used in this study.

<p>(<b>a</b>) DAADs are taken up by bacteria where they compete with endogenous D-Ala-D-Ala (DA—DA) for incorporation into PG. De novo synthesis of DA—DA is inhibited by D-cyloserine. The pentapeptide PG subunit is then flipped across the inner membrane into the periplasm where it is transglycosylated to form glycan polymers (nascent PG) and crosslinked by penicillin binding proteins (PBPs). Transpeptidation causes cleavage of the terminal D-Ala at position 5. Because the N-terminally labeled portion of DAAD becomes the amino acid at position 4 of the pentapeptide, the label is resistant to this processing and remains on the stem peptide. (<b>b</b>) PG-labeling reagents used in this study. Clickable DAADs EDA—DA and ADA—DA. Star represents the clickable amino acid.</p