Marine Tubeworm Metamorphosis Induced by Arrays of Bacterial
Phage Tail–Like Structures
Nicholas J. Shikuma
1,*
,
Martin Pilhofer
1,2,*
,
Gregor L. Weiss
1
,
Michael G. Hadfield
3,†
,
Grant
J. Jensen
1,2,†
, and
Dianne K. Newman
1,2,†
1
Division of Biology, California Institute of Technology, Pasadena, CA 91125, USA.
2
Howard Hughes Medical Institute, Pasadena, CA 91125, USA.
3
Kewalo Marine Laboratory, University of Hawai‘i at Mānoa, Honolulu, HI 96813, USA.
Abstract
Many benthic marine animal populations are established and maintained by free-swimming larvae
that recognize cues from surface-bound bacteria to settle and metamorphose. Larvae of the
tubeworm
Hydroides elegans
, an important biofouling agent, require contact with surface-bound
bacteria to undergo metamorphosis; however, the mechanisms that underpin this microbially
mediated developmental transition have been enigmatic. Here, we show that a marine bacterium,
Pseudoalteromonas luteoviolacea
, produces arrays of phage tail–like structures that trigger
metamorphosis of
H. elegans
. These arrays comprise about 100 contractile structures with
outward-facing baseplates, linked by tail fibers and a dynamic hexagonal net. Not only do these
arrays suggest a novel form of bacterium-animal interaction, they provide an entry point to
understanding how marine biofilms can trigger animal development.
Environmentally selective settlement of swimming larvae that are the propagules of most
benthic invertebrate species is a critical life-cycle stage achieved by recognizing specific
physicochemical cues (
1
,
2
). This process is of fundamental importance to the fields of
developmental biology and marine benthic community ecology—for example, the
recruitment of new larval animals is essential to sustain and disperse coral reef populations
(
1
). Economically, larval settlement is necessary for the supply of products for fisheries and
aquaculture industries worldwide (
3
) and is responsible for millions of dollars of increased
fuel consumption per year due to the biofouling of ships (
4
). Bacteria resident in surface
biofilms are now recognized as the sources of metamorphosis-inducing cues for many
invertebrates from most phyla (
2
). Indeed, the importance of microbes to the development
†
Corresponding author. hadfield@hawaii.edu (M.G.H.); jensen@caltech.edu (G.J.J.); dkn@caltech.edu (D.K.N.).
*
These authors contributed equally to this work.
Author contributions: All authors designed research. N.J.S., M.P. and G.L.W. performed research. All authors wrote the paper.
Supplementary Materials
www.sciencemag.org/content/343/6170/529/suppl/DC1
Materials and Methods
Figs. S1 to S10
Tables S1 to S4
References (
30–46
)
Movies S1 to S8
HHS Public Access
Author manuscript
Science
. Author manuscript; available in PMC 2016 July 18.
Published in final edited form as:
Science
. 2014 January 31; 343(6170): 529–533. doi:10.1126/science.1246794.
Author Manuscript
Author Manuscript
Author Manuscript
Author Manuscript
and health of diverse animals is becoming increasingly appreciated (
5
). Yet our
understanding of how these microbes interact with their hosts is only in its infancy.
The relation between the marine tubeworm
Hydroides elegans
and the bacterium
Pseudoalteromonas luteoviolacea
is a model for the study of invertebrate metamorphosis (
2
,
6
,
7
). Bacteria from the genus
Pseudoalteromonas
are commonly isolated from marine water,
sediment, biofilms, or marine eukaryotes (
8
,
9
).
P. luteoviolacea
strain HI1, used in this
study, was isolated from a marine biofilm (
8
). Recently, Huang
et al
. (
7
) identified a set of
genes in
P. luteoviolacea
whose products are essential to metamorphosis of
H. elegans
.
However, the specific cue that triggers this bacterium-mediated developmental transition
remained unknown.
In the vicinity of the
P. luteoviolacea
genes identified as essential to the induction of
H.
elegans
metamorphosis (
7
) (Fig. 1, A and B), we identified a cluster of open reading frames
(ORFs) predicted to encode components of phage tail–like structures, known as bacteriocins
(fig. S1). Bacteria typically use bacteriocins to kill other bacteria by puncturing their
membrane, causing depolarization (
10
,
11
). R-type bacteriocins resemble contractile phage
tails, similar to type VI secretion systems (T6SS) of Gram-negative bacteria (
12
). Phage
tail–like bacteriocins have a contractile sheath, inner tube, baseplate components and tail
fibers, but lack a DNA-filled head and are therefore not replicative. Bacteriocin-like
structures can mediate several bacterial pathogen-animal interactions, for example, by
causing antifeeding activity in grass grubs (
13
) and insecticidal activity against wax moths
(
14
). No phage tail–like structures are currently known to mediate an interaction that is
beneficial for the animal. On the basis of their predicted role in inducing metamorphosis, we
named the ORFs surrounding those identified by Huang
et al
. (
7
) the metamorphosis-
associated contractile structure (
mac
) genes.
To determine whether the
mac
genes play a role in tubeworm metamorphosis, we made in-
frame deletions of genes encoding putative sheath (
macS
), tube (
macT1
and
macT2
), and
baseplate (
macB
) [previously identified by Huang
et al
. (
7
)] proteins (fig. S1). These
deletion strains grew identically in rich medium (Fig. 1C) but were unable to induce
metamorphosis (Fig. 1D). Complementation of mutant strains in trans with
mac
genes
resulted only in modest restoration of metamorphosis (fig. S2A). When the
mac
genes were
replaced in their native chromosomal loci, metamorphosis induction was restored (fig. S2B).
In addition to the
mac
gene cluster, we identified a second phage tail–like bacteriocin locus
(bacteriocin-2) containing two genes predicted to encode putative tube and sheath proteins
and a third gene cluster predicted to encode proteins similar to tube (Hcp) and sheath
proteins (VipA/B) from a T6SS (table S1). In contrast to the Δ
mac
mutants, strains
containing in-frame deletions of genes predicted to encode tube and sheath proteins of
bacteriocin-2 (Δ
bact2
) or T6SS (Δ
vipABhcp
) still induced metamorphosis of
H. elegans
similarly to wild type (Fig. 1D).
To test whether the
mac
gene-cluster is responsible for producing phage tail–like structures,
we compared negatively stained electron micrographs of cultures of
P. luteoviolacea
wild
type—producing MAC, bacteriocin-2, and T6SS—and mutants producing only MACs
(Δ
vipABhcp
Δ
bact2
), producing only bacteriocin-2 (Δ
vipABhcp
Δ
macS
Δ
macB
), or lacking
Shikuma et al.
Page 2
Science
. Author manuscript; available in PMC 2016 July 18.
Author Manuscript
Author Manuscript
Author Manuscript
Author Manuscript
T6SS, bacteriocin-2, and MACs (Δ
vipABhcp
Δ
bact2
Δ
macS
Δ
macB
). Contracted and
disassembled phage tail−like bacteriocins, as well as aggregated sheaths, were observed in
the extracellular space of wild-type cells (Fig. 2A). Cultures producing only MACs
(Δ
vipABhcp
Δ
bact2
) contained dense aggregates of contracted sheaths (length 135 ± 4 nm,
n
= 13) and possibly tube structures (Fig. 2B), whereas cultures producing only bacteriocin-2
(Δ
vipABhcp
Δ
macS
Δ
macB
) contained contracted and disassembled individual phage tail–
like structures, with shorter contracted sheaths (94 ± 3 nm,
n
= 13) (Fig. 2C). No sheath or
phage tail–like structures were detected in the strain lacking MACs, bacteriocin-2, and T6SS
(Δ
vipABhcp
Δ
bact2
Δ
macS
Δ
macB
) (Fig. 2D). Similarly, we observed sheaths or bacteriocins
in purifications from the same strains, except that dense aggregates of MACs were not
observed (fig. S3, A to D). Mass spectrometry of two bands present in an SDS–
polyacrylamide gel electrophoresis (SDS-PAGE) gel of purified proteins from the strain
producing only MAC (Δ
vipABhcp
Δ
bact2
)—not seen in the sample from the strain lacking
MAC, bacteriocin-2, and T6SS (Δ
vipABhcp
Δ
bact2
Δ
macS
Δ
macB
) (fig. S4)—revealed three
peptides matching the putative MAC sheath protein (MacS) and one peptide matching a
putative MAC tube protein (MacT2).
To determine the phylogenetic placement of MACs, we compared sequences of domains
from MacT1 andMacT2 to similar domains from the phage T4 Gp19 tail tube (PF06841)
protein family (e-value = 2.1e-32 and 3.1e-13, respectively). MacT1 and MacT2 grouped
with bacterial Gp19 proteins within distinct clades (Fig. 1E) and are closely related to the
phage tail–like bacteriocin tube proteins from
Serratia entomophila
(
13
) and
Photorhabdus
asymbiotica
(
14
). Comparative genomics also revealed that multiple genes in the
P.
luteoviolacea
gene cluster are homologous to genes from the
S. entomophila afp
locus, with
some conservation of synteny (fig. S1, table S1), which suggested a common evolutionary
origin. It is noteworthy that MacT1 and MacT2 are related to tube proteins predicted to
function as part of T6SSs (
15
) in the parasitic wasp symbiont,
Cardinium hertigii
(
16
), and
the amoeba symbiont,
Candidatus Amoebophilus asiaticus
(
17
) (Fig. 1E).
Given the relatedness of MACs to phage and phage tail–like structures, we investigated
whether
P. luteoviolacea
produces and releases MACs extracellularly. To track MAC
localization in vivo, we constructed a strain encoding a C-terminal fusion of MacB with
superfolder green fluorescent protein (sfGFP) (
18
). When the native chromosomal
macB
gene was replaced with one encoding MacB-sfGFP, the recombinant strain induced
metamorphosis to levels comparable with that of wild type (Fig. 1D). Fluorescence light
microscopy revealed that the MacB-sfGFP fusion protein localized extracellularly when
broth cultures reached stationary phase and produced approximately 0.5- to 1.0-μm-wide
ringlike signals, whereas the untagged strain showed no fluorescence (Fig. 2, E and F).
MacB-sfGFP expression was observed by using three different marine media [ASWT,
NSWT, and 2216 (fig. S5)], which suggests that extracellular release is dependent neither on
soluble factors present in natural seawater nor on a specific nutrient-rich medium. Of the
cells in a population, 2.4% (
n
= 1244) showed intra-cellular GFP expression, many of them
seemingly in the process of cell lysis. Time-lapse microscopy revealed that lysis of a subset
of cells precedes the appearance of extracellular MacB-sfGFP (movie S1).
Shikuma et al.
Page 3
Science
. Author manuscript; available in PMC 2016 July 18.
Author Manuscript
Author Manuscript
Author Manuscript
Author Manuscript
Electron cryomicroscopy (
19
) of a strain producing only MACs (Δ
vipABhcp
Δ
bact2
)
allowed us to visualize these structures at high resolution in a near-native state. Four out of
162 cells (2.5%) showed intracellular phage tail–like bacteriocins. This percentage of MAC-
containing cells matches the percentage of cells harboring MacB-sfGFP. Electron
cryotomography (ECT) of these four cells revealed that the entire cytoplasm was packed
with clusters of MACs, which in some cases appeared to be connected by a filamentous
mesh (white arrow in Fig. 3A, movie S2). The shape and membrane structures of MAC-
producing cells indicated that they were about to undergo lysis (fig. S6).
We then asked whether enriched MAC preparations were sufficient to induce metamorphosis
in the absence of bacterial cells. MAC preparations obtained by standard bacteriocin
purification protocols (
12
) failed to induce
H. elegans
metamorphosis. MACs likely contract
upon purification, and the arrays fall apart, consistent with our observation that MACs were
almost exclusively in a contracted state when visualized by negative-stain electron
microscopy (EM). A gentle purification from wild-type
P. luteoviolacea
resulted in a MAC
preparation that induced metamorphosis of
H. elegans
(Fig. 2G), whereas extracts from a
MAC mutant (Δ
macB
) did not. Assays were performed with extract concentrations derived
from the equivalent of 10
7
cells/ml (100× dilution). Filtering extracts from wild-type cells
through a 0.45-μm filter abolished the metamorphic effect, consistent with the observation
that MacB-sfGFP forms >0.45-μm aggregates. Concentrated bacteriocin extracts (derived
from the equivalent of 10
8
cells/ml, 10× dilution) caused 100% larval death after 24 hours,
which indicated that MACs or copurifying constituents can have toxic effects at high doses.
We do not know the concentration of MACs in laboratory or natural marine biofilms.
To address whether another factor present in the MAC lysate was responsible for inducing
metamorphosis independent of MACs, we constructed
macB-sfgfp
strains in mutants
lacking the MAC sheath protein (Δ
macS
), or MAC tube proteins (Δ
macT1
or Δ
macT2
). We
observed extracellular MacB-sfGFP fluorescence in all mutants (fig. S7, C to E). Levels of
MacB-sfGFP were comparable between a
macB-sfgfp
strain and strains with deletions of
macS, macT1
, or
macT2
, as determined by immunoblot analysis with antibodies against
GFP (fig. S7, F and G), which suggested that the expression and stability of MacB-sfGFP is
not dependent on other MAC components. These results indicate that other biomolecules
derived from lysed cells are insufficient to promote metamorphosis of
H. elegans
in the
absence of functional MACs. Although MACs are necessary for metamorphosis, whether
they are sufficient remains to be determined. Additional ORFs in the
mac
gene cluster, such
as those identified by Huang
et al
. (
7
) (i.e., ORF2, ORF3, and ORF4A/B/C) (fig. S1), might
also contribute to the MAC structure or the direct induction of metamorphosis.
We used the same MAC extracts shown to promote metamorphosis of
H. elegans
in
bacteriocidal activity assays with closely related
P. luteoviolacea
strains and species (
P.
luteoviolacea
strain ATCC 33492,
P. piscicida
, and
P. haloplanktis
). These MAC extracts did
not kill close relatives of
P. luteoviolacea
HI1 (fig. S8), unlike bacteriocins produced by
other bacteria (
20
,
21
). It remains to be determined whether these or other types of MACs
can kill bacterial species under different conditions.
Shikuma et al.
Page 4
Science
. Author manuscript; available in PMC 2016 July 18.
Author Manuscript
Author Manuscript
Author Manuscript
Author Manuscript
We characterized MAC aggregates using ECT to compare their structure with that of other
phage tail–like bacteriocins. In a frozen, near-native state, we observed MACs forming
arrays of multiple contractile phage tail–like structures (Fig. 3 and movies S3 and S4). A
typical MAC array contained close to 100 individual phagelike tails (96 for the one shown in
Fig. 3, B to I), with dimensions matching the size of MacB-sfGFP fluorescence (fig. S9 and
table S2). Intracellular MACs were more tightly packed than their extracellular counterparts
(table S2), which suggested that MAC arrays expand upon cell lysis. MACs radiate into a
hemisphere, originating from an amorphous center (yellow arrow in Fig. 3B). Distal MAC
ends were arranged into a regular array, surrounded by a filamentous hexagonal net (white
arrow in Fig. 3E). MACs were oriented with their baseplates (“B” in Fig. 4B) outward, with
filamentous structures (probably tail fibers; orange arrows in Fig. 3F) emanating from the
baseplates and appearing to connect individual MACs to each other (“TF” in Fig. 4B; see
also movie S3). To our knowledge, such ordered multitailed arrays have not previously been
observed for other types of phagelike structures.
Individual MACs resembled contractile phage tail–like structures (Fig. 4). Sheaths were
observed in extended (“E” in Fig. 4B, table S2) and contracted (“C” in Fig. 4B, table S2)
forms, with homogeneous and helical surface patterns, respectively. In some cases, the inner
tube (“T” in Fig. 4, A and B, and table S2) was observed. We classified the observed MAC
conformations into “tube only” (T), “extended” (E), “contracted with jammed tube” (J),
“contracted with fired tube” (C) and “contracted without tube” (S) (Fig. 4, A to E, and
movie S5). The “extended” conformation represented 73% of MACs within the aggregate
shown (Fig. 3). These different structural classes likely represent different states in a
functional sequence. In analogy to phage tail–like structures, the “tube only” state may be a
partially assembled MAC, whereas the “extended” MAC is in a ready-to-fire configuration
(
12
,
22
). “Contracted with fired tube” and “contracted without tube” are likely MACs that
have fired. “Contracted with jammed tube” could represent fired MACs that failed to propel
the tube. In this conformation, the sheath is contracted as indicated by the helical ridges
[assembling T6SS sheaths are in the extended conformation (
12
)], and the sheath length of 5
of 7 structures matches the length of a fired MAC (table S2).
We averaged 20 and 25 subtomograms of MACs in the “extended” (Fig. 4P, movie S6) and
“contracted without tube” (Fig. 4Q, movie S7) states, respectively. Averages of
subtomograms show the different sheath diameters, helical surface ridges on contracted
sheath (ridges indicated in Fig. 4K), baseplate symmetry and tail fibers in longitudinal (Fig.
4, J and O) and cross-sectional views (Fig. 4, F to I and K to N). In both the extended and
contracted forms twelve fibers emerge from the baseplate, cross paths, and separate to meet
at the ring-shaped vertices of the hexagonal net surrounding individual MACs (Fig. 4R,
movie S8). We speculate that six of the tail fibers originate from a single MAC, with the
remaining six fibers stemming from neighboring MACs to connect the array (Fig. 4, P to S,
orange). This six-tail fiber per MAC model (Fig. 4S) is supported by the fact that the two
arms of a phage tail fiber have a length ratio of 1:1 (
23
) and that the length is similar to the
tail fiber connections in MACs (fig. S10). The model also predicts the presence of an as-yet-
unidentified protein that forms the hexagonal net. A set of six tail pins (Fig. 4P, red) face
outwards. Because the tail pins are the most distal structure in the arrays, they are likely the
first structure to engage and sense MAC targets.
Shikuma et al.
Page 5
Science
. Author manuscript; available in PMC 2016 July 18.
Author Manuscript
Author Manuscript
Author Manuscript
Author Manuscript
We have shown that an ordered array of contractile phage tail–like structures produced by an
environmentally occurring bacterium induces metamorphosis of a marine invertebrate larva.
This discovery begins to explain how marine biofilms can trigger metamorphosis of benthic
animals. Our data suggest that MAC arrays are synthesized intracellularly by
P.
luteoviolacea
, released by cell lysis, and expand extracellularly into an ordered multi-MAC
array. How these arrays engage with larvae of
H. elegans
is an open question. In the arrays
imaged, all contracted MACs were clustered together, which suggests that their linkages
might support cooperative firing. Array formation might also multiply the total payload
delivered per interaction or favor specific engagement sites and/or geometries with MAC
targets. The evolutionary pressure to produce MACs is probably strong, given that MAC
production leads to the lysis and death of a subpopulation of cells. Whether this represents
an instance of altruistic behavior that facilitates group selection, or a neutral lytic event with
a set frequency remains to be determined. Although MAC production is beneficial for
tubeworm larvae by inducing metamorphosis, it is currently unclear how larval settlement
and metamorphosis might benefit the bacterium. It is equally possible that MACs evolved
for a completely different purpose. Note that
P. luteoviolacea
has been found to induce the
metamorphosis of coral and sea-urchin larvae (
24
,
25
). Other bacterial species also induce
metamorphosis of
H. elegans
larvae (
8
,
26
,
27
), and
mac
-like gene clusters have been
identified in the genomes of other marine bacteria (
28
). Future research into how MACs
interact with larvae might yield new insights into the mechanisms underpinning marine
animal development and ecology, with potentially important practical applications for
aquaculture and biofouling.
Supplementary Material
Refer to Web version on PubMed Central for supplementary material.
Acknowledgments
We thank B. Pernet for help with locating and identifying tubeworms and for giving us the algal strain used in this
work; A. McDowall for help with EM; Y. Huang, who created the Str
R
-strain (
7
); A. Asahina and S. Wilbur for
laboratory assistance; J. Levine for help with time-lapse microscopy; J. Ricci for help with phylogenetic analyses;
and members of the Newman group for discussions and comments on the manuscript. The Howard Hughes Medical
Institute, Z. Yu, and J. de la Cruz are acknowledged for providing access to the FEI Titan Krios at Janelia Farm and
support in data collection. N.J.S. was supported by a California Institute of Technology (Caltech) Division of
Biology Postdoctoral Fellowship. This collaboration was supported by the Caltech Center for Environmental
Microbiology Interactions, the Howard Hughes Medical Institute (D.K.N. and G.J.J.), Office of Naval Research
grants N00014-08-1-0413 and N00014-05-1-0579 (M.G.H.), NIH grant GM094800B (G.J.J.), and a gift from the
Gordon and Betty Moore Foundation (Caltech). D.K.N. and G.J.J. are Investigators of the Howard Hughes Medical
Institute. Strains obtained from the American Type Culture Collection listed in table S2 (ATCC 33492, ATCC
14393, ATCC 15057). DNA sequences encoding for
mac
, T6SS, and bacteriocin-2 genes are deposited under
GenBank accession numbers KF724687, KF724688, and KF724689, respectively. Subtomogram averages were
deposited in the Electron Microscopy Data Bank (accession numbers EMD-2543, EMD-2544, and EMD-2545).
References and Notes
1. Hadfield, MG.; Paul, VJ. Marine Chemical Ecology. McClintock, JB.; Baker, BJ., editors. Boca
Raton, FL: CRC Press; 2001. p. 431
2. Hadfield MG. Annu. Rev. Mar. Sci. 2011; 3:453–470.
3. Food and Agriculture Organization of the United Nations. The State of World Fisheries and
Aquaculture 2012. Rome: FAO; 2012.
Shikuma et al.
Page 6
Science
. Author manuscript; available in PMC 2016 July 18.
Author Manuscript
Author Manuscript
Author Manuscript
Author Manuscript
4. Schultz MP, Bendick JA, Holm ER, Hertel WM. Biofouling. 2011; 27:87–98. [PubMed: 21161774]
5. McFall-Ngai M, et al. Proc. Natl. Acad. Sci. U.S.A. 2013; 110:3229–3236. [PubMed: 23391737]
6. Nedved BT, Hadfield MG. Marine Indust.l Biofoul. 2009; 4:203–217.
7. Huang Y, Callahan S, Hadfield MG. Sci. Rep. 2012; 2:228. [PubMed: 22355742]
8. Huang SY, Hadfield MG. Mar. Ecol. Prog. Ser. 2003; 260:161–172.
9. Holmström C, Kjelleberg S. FEMS Microbiol. Ecol. 1999; 30:285–293. [PubMed: 10568837]
10. Michel-Briand Y, Baysse C. Biochimie. 2002; 84:499–510. [PubMed: 12423794]
11. Uratani Y, Hoshino T. J. Bacteriol. 1984; 157:632–636. [PubMed: 6420392]
12. Basler M, Pilhofer M, Henderson GP, Jensen GJ, Mekalanos JJ. Nature. 2012; 483:182–186.
[PubMed: 22367545]
13. Hurst MR, Glare TR, Jackson TA. J. Bacteriol. 2004; 186:5116–5128. [PubMed: 15262948]
14. Yang G, Dowling AJ, Gerike U, ffrench-Constant RH, Waterfield NR. J. Bacteriol. 2006;
188:2254–2261. [PubMed: 16513755]
15. Pukatzki S, et al. Proc. Natl. Acad. Sci. U.S.A. 2006; 103:1528–1533. [PubMed: 16432199]
16. Penz T, et al. PLOS Genet. 2012; 8:e1003012. [PubMed: 23133394]
17. Penz T, Horn M, Schmitz-Esser S. Virulence. 2010; 1:541–545. [PubMed: 21178499]
18. Pédelacq JD, Cabantous S, Tran T, Terwilliger TC, Waldo GS. Nat. Biotechnol. 2006; 24:79–88.
[PubMed: 16369541]
19. Pilhofer M, Ladinsky MS, McDowall AW, Jensen GJ. Methods Cell Biol. 2010; 96:21–45.
[PubMed: 20869517]
20. Köhler T, Donner V, van Delden C. J. Bacteriol. 2010; 192:1921–1928. [PubMed: 20118263]
21. Gebhart D, et al. J. Bacteriol. 2012; 194:6240–6247. [PubMed: 22984261]
22. Leiman PG, et al. Virol. J. 2010; 7:355. [PubMed: 21129200]
23. Cerritelli ME, Wall JS, Simon MN, Conway JF, Steven AC. J. Mol. Biol. 1996; 260:767–780.
[PubMed: 8709154]
24. Tran C, Hadfield MG. Mar. Ecol. Prog. Ser. 2011; 433:85–96.
25. Huggett MJ, Williamson JE, de Nys R, Kjelleberg S, Steinberg PD. Oecologia. 2006; 149:604–
619. [PubMed: 16794830]
26. Unabia CRC, Hadfield MG. Mar. Biol. 1999; 133:55–64.
27. Lau SC, Mak KK, Chen F, Qian P-Y. Mar. Ecol. Prog. Ser. 2002; 226:301–310.
28. Persson OP, et al. Environ. Microbiol. 2009; 11:1348–1357. [PubMed: 19207573]
29. Anisimova M, Gascuel O. Syst. Biol. 2006; 55:539–552. [PubMed: 16785212]
Shikuma et al.
Page 7
Science
. Author manuscript; available in PMC 2016 July 18.
Author Manuscript
Author Manuscript
Author Manuscript
Author Manuscript
Fig. 1.
P. luteoviolacea mac
genes are required for metamorphosis of
H. elegans
and are similar to
genes encoding phage tail–like structures
(
A
and
B
) Metamorphically competent
H. elegans
larva (A) and juvenile adult (B) 12 hours
after exposure to a
P. luteoviolacea
biofilm. Scale bars, 50 μm. (
C
) Growth of
P.
luteoviolacea
wild type and mutants containing in-frame deletions of
mac
genes. OD, optical
density, a measure of absorbance. (
D
) Metamorphosis (%) of
H. elegans
in response to
biofilms of
P. luteoviolacea
wild type; and Δ
macB
, Δ
macT1
, Δ
macT2
, Δ
macS
, Δ
bact2
, and
Δ
vipABhcp
mutants; and
macB-sfgfp
fusion strains. Sterile artificial seawater (no bacteria)
was used as a negative control. Error bars represent standard deviations (
n
= 4). (
E
)
Maximum likelihood unrooted phylogeny of the Gp19 protein family. Gp19-like protein
domains originating from bacteria and phages are highlighted in blue and yellow,
respectively. Nodes with approximate likelihood-ratio test (aLRT) (
29
) values ≥0.8 are
marked with a black circle. Scale bar indicates amino acid substitutions per site.
Shikuma et al.
Page 8
Science
. Author manuscript; available in PMC 2016 July 18.
Author Manuscript
Author Manuscript
Author Manuscript
Author Manuscript
Fig. 2. MACs are phage tail–like structures, are released by cell lysis, and mediate
metamorphosis of
H. elegans
(
A
to
D
) Negative stain EM of
P. luteoviolacea
(A) wild type, (B) Δ
vipABhcp
Δ
bact2
, (C)
Δ
vipABhcp
Δ
macS
Δ
macB
, and (D) Δ
vipABhcp
Δ
bact2
Δ
macS
Δ
macB
. Aggregated sheaths
are indicated by an arrow in (B). Scale bars, 200 nm. (
E
and
F
) Micrographs of merged
phase-contrast and fluorescence images of
P. luteoviolacea
(E) wild-type and (F)
macB-sfgfp
strains (see movie S1). Fluorescence of MacB-sfGFP is shown in green. Scale bar, 5 μm. (
G
)
Metamorphosis (%) of
H. elegans
in response to cell-free extracts from
P. luteoviolacea
wild
type and Δ
macB
mutant. Extracts unfiltered and 0.45-μm filtered are indicated. Sterile
artificial seawater (no bacteria) was used as a negative control. Error bars represent standard
deviation (
n
= 4).
Shikuma et al.
Page 9
Science
. Author manuscript; available in PMC 2016 July 18.
Author Manuscript
Author Manuscript
Author Manuscript
Author Manuscript
Fig. 3. MACs are assembled intracellularly and expand as an ordered array upon cell lysis
P. luteoviolacea
Δ
vipABhcp
Δ
bact2
mutant cells and extracellular aggregates were imaged by
ECT (shown are 16.8-nm-thick slices). (
A
) The cytoplasm of a subset of cells (4 in 162) was
packed with MAC aggregates, and cells appeared in the process of lysis (see movie S2). I,
inner membrane; O, outer membrane; S, storage granule; white arrow, filamentous
connections. (
B
to
G
) Extracellular MAC arrays are highly ordered (shown are 2D slices
through a tomogram at different z-heights) (see movies S3 and S4). Yellow arrow,
amorphous core; green arrow, inner tube; blue arrow, sheath; white arrow, filamentous
hexagonal net; orange arrow, tail fibers; P, presumably polymerized sheath protein. (
H
and
I
)
MAC arrays were hemispherical with MACs coalescing in an amorphous core and the
baseplates hexagonally arranged on the surface. Individual MACs were connected by tail
fibers and surrounded by a hexagonal net. Different views of a segmented model of the array
are shown. Slice z-heights in (B) to (G) are indicated with the corresponding panel letter.
Scale bars, 100 nm.
Shikuma et al.
Page 10
Science
. Author manuscript; available in PMC 2016 July 18.
Author Manuscript
Author Manuscript
Author Manuscript
Author Manuscript
Fig. 4. MACs are observed in different functional states and connected by tail fibers
(
A
) Side view of a MAC in the tube only T state. (
B
) Side views of MACs in extended, E,
and contracted, C, states. B, baseplate; TF, tail fibers; T, inner tube. (
C
) Side view of a MAC
in the contracted sheath without tube S state. (
D
) Side view of a MAC in the contracted state
with jammed tube J. Scale bars in (A) to (D), 100 nm. Tomographic slices that are 16.8 nm
thick are shown in (A) to (D). (
E
) Schematic of different functional states. Numbers indicate
the quantity of each state found in the MAC array shown in Fig. 3, (B) to (I). (
F
to
J
)
Subtomogram average of extended MACs. Cross-sectional slices at different z-heights are
shown in (F) to (I) and their positions are indicated in the side view (J). Slices that are 8.4
nm thick are shown. Scale bars, 10 nm. (
K
to
O
) Subtomogram average of contracted
MACs. Cross-sectional slices at different z-heights are shown in (K) to (N), and their
Shikuma et al.
Page 11
Science
. Author manuscript; available in PMC 2016 July 18.
Author Manuscript
Author Manuscript
Author Manuscript
Author Manuscript
positions are indicated in the side view (O). Slices that are 8.4 nm thick are shown. Scale
bars, 10 nm. (
P
to
R
) Isosurface of subtomogram averages (see movies S6 to S8) of an
extended (P) and a contracted (Q) MAC and of the tail fiber junction (R). Tail pins and
baseplate, red; tail fibers, orange; inner tube and spike, green; sheath, blue; hexagonal net,
white. (
S
) Schematic model of tail fiber connections in the MAC array. Each MAC unit
contributes six tail fibers that connect to neighboring MAC units. The central MAC unit is
colored in the color code used in (P) to (R). A second MAC unit is colored magenta.
Shikuma et al.
Page 12
Science
. Author manuscript; available in PMC 2016 July 18.
Author Manuscript
Author Manuscript
Author Manuscript
Author Manuscript