The Dynamically Evolving Nematocyst Content of an
Anthozoan, a Scyphozoan, and a Hydrozoan
Tamar Rachamim,y,1 David Morgenstern,y,2 Dikla Aharonovich,1 Vera Brekhman,1 Tamar Lotan,*,1 and
Daniel Sher*,1
1
Department of Marine Biology, Leon H. Charney School of Marine Sciences, University of Haifa, Haifa, Israel
Proteomics Resource Center, Langone Medical Center, New York University
yThese authors contributed equally to this work.
*Corresponding author: E-mail: [email protected]; [email protected].
Associate editor: Willie Swanson
2
Abstract
Nematocytes, the stinging cells of cnidarians, are the most evolutionarily ancient venom apparatus. These nanosyringelike weaponry systems reach pressures of approximately 150 atmospheres before discharging and punching through the
outer layer of the prey or predator at accelerations of more than 5 million g, making them one of the fastest biomechanical events known. To gain better understanding of the function of the complex, phylum-specific nematocyst
organelle, and its venom payload, we compared the soluble nematocyst’s proteome from the sea anemone Anemonia
viridis, the jellyfish Aurelia aurita, and the hydrozoan Hydra magnipapillata, each belonging to one of the three basal
cnidarian lineages which diverged over 600 Ma. Although the basic morphological and functional characteristics of the
nematocysts of the three organisms are similar, out of hundreds of proteins identified in each organism, only six are
shared. These include structural proteins, a chaperone which may help maintain venon activity over extended periods,
and dickkopf, an enigmatic Wnt ligand which may also serve as a toxin. Nevertheless, many protein domains are shared
between the three organisms’ nematocyst content suggesting common proteome functionalities. The venoms of Hydra
and Aurelia appear to be functionally similar and composed mainly of cytotoxins and enzymes, whereas the venom of the
Anemonia is markedly unique and based on peptide neurotoxins. Cnidarian venoms show evidence for functional
recruitment, yet evidence for diversification through positive selection, common to other venoms, is lacking. The final
injected nematocyst payload comprises a mixture of dynamically evolving proteins involved in the development, maturation, maintenance, and discharge of the nematocysts, which is unique to each organism and potentially to each
nematocyst type.
Key words: nematocyst, cnidaria, toxin, mass spectrometry, transcriptome, anemone, jellyfish, hydra, venom, protein
domain.
Article
Introduction
Cnidarians, such as sea anemones, corals, jellyfish and hydra,
are one of the most ancient and morphologically simple
animal phyla. From their origins at least 700 Ma during the
Precambrian period, these soft-bodied, mostly sessile organisms have seen the emergence (and decline) of countless new
life-forms with more complex modes of environmental sensing and information processing, locomotion, and feeding
(Cartwright et al. 2007; Shinzato et al. 2011). Yet cnidarians
continue to flourish in all aquatic environments, defending
themselves from, and even preying upon, more active, motile
and complex organisms. A significant part of the success of
cnidarians can be attributed to their ability to produce a wide
array of toxins and other bioactive molecules for defense and
prey capture (Anderluh and Macek 2002; Norton 2009; Sher
and Zlotkin 2009). It can also be attributed to their ability to
deliver some of their chemical arsenal into their prey or predators using a complex and elaborate venom delivery system—
the stinging cells, or nematocytes (synonym cnidocytes; Lotan
et al. 1995; Tardent 1995; Nuchter et al. 2006; Beckmann and
Ozbek 2012).
Nematocytes are a defining feature of cnidarians, observed
in fossils from the Middle Cambrian period, approximately
505 Ma (Cartwright et al. 2007; Young and Hagadorn 2010).
Most of the nematocyte cell volume is taken up by the
nematocyst—a large complex, Golgi-derived secretory vesicle
(Tardent 1995; Engel et al. 2002; David et al. 2008).
Nematocytes undergo a process of assembly, maturation,
and migration, at the end of which they are mounted
where they will be deployed—typically the ectoderm of the
hunting tentacles (Denker et al. 2008). Once fully assembled,
the nematocytes become a loaded microweaponry system
capable of punching through the integument of prey or predator at accelerations of more than approximately 5 million g,
inserting a thin needle-like tubule through which venom is
injected (Holstein and Tardent 1984; Nuchter et al. 2006). The
delivered venom induces rapid paralysis for prey capture and
may cause pain, possibly for predator deterrence (Brodie
2009). Cnidarian nematocyst venoms exhibit additional pharmacological and physiological activities such as cytolysis, dermatonecrosis, proteolysis, and vasopermeation (Sher et al.
2005, and references therein). How the different venom
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Mol. Biol. Evol. 32(3):740–753 doi:10.1093/molbev/msu335 Advance Access publication December 16, 2014
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Nematocyst Content . doi:10.1093/molbev/msu335
toxins work together to produce the systemic effect of prey
paralysis or predator deterrence is still not well understood.
The structure of the nematocyst capsule itself is composed
of a mesh of proteins cross-linked by intermolecular disulfide
bridges providing strength and elasticity, with additional
structural proteins forming the tubule and its barbs (reviewed
by Beckmann and Ozbek [2012]). The highly packed matrix
inside the capsule comprised mainly poly--glutamate associated with cations. Upon nematocyte activation, water flows
through the nematocyst wall causing the disassociation of the
aggregate matrix and resulting in an increase in osmotic pressure which leads to the explosive nematocyst discharge
(Weber 1989, 1990; Szczepanek et al. 2002; Ayalon et al.
2011). Once the nematocyst has discharged, the venom is
injected through the tubule and into the prey (Lotan et al.
1996; Basulto et al. 2006).
Recently, several proteomic studies have shown that nematocysts from various cnidarians contain tens to hundreds of
proteins, many of which are unique to cnidarians and of
which very few have been characterized (Balasubramanian
et al. 2012; Brinkman et al. 2012; Li et al. 2012; Moran et al.
2013; Weston et al. 2013). This surprising complexity raises
several questions: What are the roles of these proteins? Are all
of them essential to the nematocyst function? Which of these
proteins is a bioactive component of the venom, inducing
paralysis, tissue damage or pain? One way of answering these
questions is to identify “core” nematocyst proteins, which, if
conserved among distantly related cnidarians, are likely to be
important for nematocyst structure or function. Here, we
compare the soluble contents of isolated nematocysts from
three different cnidarians: The Anthozoan sea anemone
Anemonia viridis (previously A. sulcata), the Schyphozoan
jellyfish Aurelia aurita, and the Hydrozoan Hydra magnipapillata (we refer to these organisms throughout the paper as
“Anemonia,” “Aurelia,” and “Hydra”). Each of these organisms
belongs to one of the three basal cnidarians lineages which
diverged over 600 Ma. Hydra has been a model organism for
over 260 years (Trembley 1744), whereas Anemonia and
Aurelia are emergent models for studying endosymbiosis
and development (e.g., Moya et al. 2012; Fuchs et al. 2014)
and the former is also a rich source of toxins and other pharmacologically active peptides (Oliveira et al. 2012). We describe commonalities and differences among the soluble
components of the nematocysts from these three organisms,
focusing on the evolutionary emergence of their functional
toxin content.
Results and Discussion
Nematocysts from Different Cnidarian Classes Share
Very Few Common Proteins
We isolated pure undischarged nematocysts from the three
cnidarians using density centrifugation (fig. 1 and Materials
and Methods), physically ruptured them without the use of
chaotrophic agents, and submitted only the water-soluble
supernatant (which likely does not include many of the structural proteins), to bottom-up mass spectrometry (MS/MS). In
parallel, we sequenced the transcriptomes of A. viridis and Au.
850-650Ma
570Ma
Hydra
magnipapillata
Aurelia
aurita
Anemonia
viridis
Proteins detected by MS/MS
291
374
737
FIG. 1. Images of Hydra, Aurelia, and Anemonia and their isolated nematocysts. The phylogeny is shown above the images. Divergence times
(Ma, millions of years ago) are according to Park et al. (2012). The
diameters of Aurelia and Anemonia are approximately 30 cm and approximately 5 cm, respectively, and the length of Hydra is approximately
2 cm. The number of MS/MS identified proteins in the nematocyst
payload of each organism is shown. The scale bar in the nematocyst
images represents 10 mm. The Aurelia and Anemonia photos were
kindly provided by Rami Tsadok and Gil Rilov, respectively.
aurita as well as, with low coverage, the genome of A. viridis
(fig. 1) (Brekhman et al. 2014). Together with the published
genome of H. magnipapillata (Chapman et al. 2010), these
data sets enabled us to identify and obtain gene sequences for
a total of approximately 290–740 proteins from the nematocysts of each organism (fig. 1, supplementary figs. S1 and S2
and Excel file S1, Supplementary Material online).
We first asked whether we can identify a set of proteins
common to the nematocysts from all three cnidarian classes.
We hypothesized that such a core proteome would be comprised proteins with an ancient and conserved role in the
nematocyst function and speculated that common toxins
may have been used by the common ancestor of the three
cnidarian lineages to prey upon early Cambrian organisms. To
our surprise, out of the hundreds of nematocyst proteins
identified by shotgun proteomics from each of the organisms,
only six proteins were common to all cnidarians (fig. 2A and
table 1), conservatively defined as either being Reciprocal Best
BLAST [Basic Local Alignment Search Tool] Hits (RBBHs)
(Altenhoff and Dessimoz 2009) or having identical annotations, with an additional quality control step of verifying a
high-level of protein identity over most of the coding sequence. This number represents the lower bracket for the
number of common proteins; however, we do not expect it
to change considerably in the future: The quantity of MS2
spectra acquired, high compatibility between transcriptome
and proteome (obtained from the same population, or, in the
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Rachamim et al. . doi:10.1093/molbev/msu335
A
Anemonia
B
Anemonia
Gamma-glutamyl transpeptidase
Protein-protein:
EGF-like, vWF,
Thrombospondin repeats
Metalloproteases
Peptidases
Protease inhibitors (Kunitz)
Peptidyl-Prolyl cis-trans
isomerase
6
220
Hydra
174
Hydra
74
158
86
65
134
Thyroglobulin type-I
Aurelia
44
69
Esterase , SGNH-type
592
21
49
Carbonic anhydrase
143
Protein-carbohydrate:
Lectins (Concavalin, type C)
Glycoside hydrolase
Aurelia
Gamma-glutamyl
hydroase
Phospholipase A2
C
Unknown or other:
MAC-PF
Histone, CAP
ShKT
HSP70
Actin
Tubulin-related
EF-hand Ca-binding
Domain abundance
Domain abundance
D
Domain rank
Domain rank
FIG. 2. Comparison of the three organism’s nematocyst proteomes. Venn diagram showing the number of proteins (A) and InterPro domains (B)
shared among the soluble nematocyst proteomes of Hydra, Anemonia, and Aurelia. A protein was defined as shared if it was an RBBH (e value cutoff
105) or had identical meaningful annotations after BLAST2GO analysis (see Materials and Methods). The domains highlighted represent some of the
domains identified (see supplementary Excel files, Supplementary Material online, for the full list). Rank-abundance curves of the InterPro domains
identified in the three organisms, colored by functional category (C) and by their presence in one or more of the organisms (D). The inset graph in panel
(C) focuses on the 50 most abundant domains for clarity, the inset in panel (D) represents the mean abundance of the same InterPro domains in three
replicate groups of 1,000 randomly selected proteins from the total Hydra proteome.
Table 1. Proteins Found in Nematocysts from All Three Organisms.
Protein
Nematocyst structure or biosynthesis
Nematogalectin
Elongation factor-1a
Multipurpose proteins
Dickkopf-3 related
Protein folding
HSP70 kDa
Unknown function
Secreted protein similar to LOC100200141
Uncharacterized 4-transmembrane domain protein
Anemonia ID
Aurelia ID
Hydra ID
Identified in
Nematocysts from
Other Organisms
FK727671_1
FK742467_1
comp195148_c1_seq1_5
comp173503_c2_seq3_4
306408168
221124734
Cf, Nv
Cf, Sm
comp19806_c0_seq1
comp187795_c0_seq1_5
22111549
Cf, Sm
FK745997_1
comp192177_c1_seq1_2
221120291
Cf a, Sm
FK748315_1
comp194496_c0_seq2_4
comp194496_c1_seq3_2
comp195147_c0_seq1_2
22110747
FK721300_1
22110228
NOTE.—Cf, Chironex fleckeri (Brinkman et al. 2012); Os, Olindias sambaquensis (Weston et al. 2013); Sm, Stomolophus meleagris (Li et al. 2012); Nv, Nematostella vectensis (Moran
et al. 2013).
a
A 78 kDa protein closely related to HSP70 was identified in C. fleckeri.
case of Anemonia, the same individual) and the use of an
error-tolerant search engine, maximized high-confidence protein identification. Notably, the fraction of nematocyst proteins shared among the three organisms (approximately 2%
of those identified in Hydra nematocysts) is much lower than
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the fraction of the total predicted Hydra proteome (comprising approximately 18,000 proteins) shared with the translated
Anemonia and Aurelia transcriptomes (RBBHs, 15.8%).
Furthermore, although Hydra have been suggested to
evolve rapidly compared with other cnidarians (Park et al.
Nematocyst Content . doi:10.1093/molbev/msu335
2012), the fraction of the nematocyst proteome shared between Hydra and the other two organisms is not smaller than
that shared between Anemonia and Aurelia (fig. 2A), suggesting that the low number of common proteins is not due to
our choice of this specific model organism. Why the nematocyst proteomes share so few proteins compared with the
full proteome, and whether this represents more rapid evolution, directional selection pressure, or some other evolutionary cause, is currently unclear. Interestingly, these
common proteins seem to largely belong to house-keeping
functions of the nematocysts rather than to the venom itself.
One of the six common proteins, Nematogalectin, is
known to be a major structural component of the nematocyst tubule (Hwang et al. 2010). A second protein, Elongation
Factor 1-alpha-like (EF-1), is a key component of the protein
translation machinery, and also has other functions, such as
mediating the dynamics of microtubule bundling and dissociation (Sasikumar et al. 2012). We hypothesize that EF-1
may bind and potentially shape the microtubule scaffold
which surrounds the developing nematocyst (Engel et al.
2002), as well as possibly take part in the localized production
of nematocyst proteins that do not contain signal sequences
and thus are not delivered through the secretory pathway
(almost one-third of the nematocyst proteome; Hwang et al.
2007; Balasubramanian et al. 2012). A third protein, dickkopf,
is a secreted antagonist of the Wnt pathway which is involved
in regeneration and axis formation, and one isoform is also
expressed in the differentiating nematocytes of Hydra
(Fedders et al. 2004; Augustin et al. 2006; Guder et al. 2006).
Intriguingly, dickkopf also reveals sequence similarity with
venom proteins, toxic skin secretions and peptides known
to interact with the voltage gated K+ channel (see supplementary fig. S3, Supplementary Material online, for details),
raising the possibility that this protein may in fact function as
a neurotoxin. The fourth protein, Heat Shock Protein 70
(HSP70), is a chaperone which promotes the correct folding
of unfolded or partially misfolded proteins. A unique aspect of
venom is that once the toxins have been produced and correctly folded, they often need to await in an active form, for
extended periods of time (days or even weeks) until the prey
or predator has been detected and the venom comes into
action. Chaperones such as HSP70, in combination with other
proteins such as disulfide isomerases (found in Hydra; supplementary Excel file S1, Supplementary Material online) and
peptidyl-prolyl cis–trans isomerases (found in all three organisms, fig. 2B), may fulfill this role, and indeed are found in the
venoms of many different organisms including wasps, snakes,
and box jellyfish (Rioux et al. 1998; Brinkman et al. 2012;
Sookrung et al. 2014). The last two proteins common to all
three organisms, one of which is a secreted protein and one of
which is predicted to have 4–5 transmembrane domains, do
not reveal similarity to any known proteins.
In addition to the proteins found in all three organisms,
three additional protein families caught our attention. The
first family, histones, is found in the nematocysts of all three
organisms, albeit with different histone combinations in each
organism (Anemonia has histone 2B and 3.2, Aurelia has histone 2A, and Hydra has histones 2A, 2B and 3). Histones are
MBE
usually associated with their role in chromatin structure, yet
as positively charged proteins either may bind the negatively
charged poly--glutamate, a major component of the nematocyst matrix, or may interact with specific proteins through
the von-Wildebrand Factor domains which are common in
the nematocyst proteome (fig. 2B) (Ward et al. 1997).
However, many free histones have strong antimicrobial and
hemolytic activities which underlie their secondary role in
innate immunity (Patat et al. 2004; Kawasaki et al. 2008;
Smith et al. 2010). Indeed, histones have recently been identified also in bee venom (Li et al. 2013). Thus, it is tempting to
speculate that the histones found in the nematocysts from
our three model organisms, as well as in nematocysts from
the jellyfish Stomolophus meleagris (Li et al. 2012) and the box
jellyfish Chironex fleckeri (Brinkman et al. 2012), may have
roles in the maintenance of the nematocyst structure, as
venom proteins, or both.
A second protein, which we identified in the nematocysts
of Hydra, is similar to Endothelin-converting Enzyme-1
(ECE-1). ECE-1 is a zinc metalloproteinase which is involved
in the processing of endothelins, potent vasoconstrictors
which also have roles in cell proliferation, differentiation,
and other developmental processes (Zhang et al. 2001;
Macours et al. 2004). In H. vulgaris, endothelins and ECEs
are involved in both muscle contraction and foot regeneration (Zhang et al. 2001); however, the ECE-1-like protein we
have identified in the nematocysts is different from the ECE
protein involved in regeneration (supplementary figs. S4 and
S5, Supplementary Material online). ECEs have recently been
identified in the venom of the fish-hunting cone snails, where
they have been proposed to increase the effectiveness of
other venom components by cleaving and activating endogenous endothelin precursors, thus reducing blood flow in the
venom injection site and increasing local venom concentration (Safavi-Hemami et al. 2013). Furthermore, many cnidarians venoms (including those that are lethal to humans)
cause severe cardiovascular effects, including hypertension,
positive inotropism, and cardiotoxicity (Lesh-Laurie et al.
1989; Ramasamy et al. 2005; Tibballs 2006; Suput 2009;
Hughes et al. 2012). Some of these effects may be caused
by endogenous endothelins or the related sarafotoxins,
potent toxins isolated from the venom of the Israeli burrowing asp Atractaspis engadensis (Kochva et al. 1993). The
ECE-1-like protein identified in Hydra has close homologs in
the transcriptomes of Aurelia and Anemonia (supplementary
figs. S4 and S5, Supplementary Material online), and a similar
protein was also identified in the venom of C. fleckeri
(Brinkman et al. 2012). We speculate, therefore, that activation of the prey’s endothelin precursors by ECEs injected
with the nematocyst venom may represent a common and
understudied aspect of some cnidarian envenomation, with
potentially important clinical implications.
Finally, we identified in Hydra and Aurelia several proteins
that may have a role in the production or degradation of the
poly--glutamate matrix, which drives the nematocyst discharge. One of these is gamma-glutamyl hydrolase (GGH),
which hydrolyzes the gamma bonds within the poly-glutamate polymer, and for which a homolog was found
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also in the Anemonia transcriptome (supplementary fig. S6,
Supplementary Material online). Weber (1994) has previously
identified GGH activity in Hydra nematocysts, showing that it
1) hydrolyzes the gamma bond in a random location within
the poly--glutamate polymer, 2) is mostly bound to the
insoluble nematocyst matrix, 3) is active under mildly acidic
conditions (pH ~ 4), and 4) is enhanced by disulfide-reducing
conditions. These results are in agreement with vertebrate
GGHs, which are thiol-activated enzymes with maximal activity at mildly acidic pH (Kao et al. 2009). Previously, it has
been proposed that the first explosive stage of nematocyst
discharge is driven by a combination of a rapid increase in
osmotic pressure due to dissociation of the poly--glutamate
from the surrounding cations and the spring-like energy
stored in the inverted tubule (Tardent 1995; Szczepanek
et al. 2002; Ozbek et al. 2009; Ayalon et al. 2011). GGH may
contribute to the increasing osmotic pressure by starting to
hydrolyze the poly--glutamate polymer into shorter chains
immediately after the activation signal is transferred into the
nematocytes and before nematocyst discharge. It may also
maintain the driving force for the second, slower stage of
tubule evertion (Holstein and Tardent 1984) and venom injection by ongoing hydrolysis of the poly--glutamate polymer that is left within the capsule, thus maintaining a
constant increase in the osmolarity of the nematocyst content that, at least partially, counteracts the loss of osmotic
pressure as poly--glutamate is washed out of the nematocyst through the tubule.
Common Domains Define a Functional Profile Shared
among Nematocysts from Different Cnidarians
Does the lack of common proteins imply that the nematocyst
payloads are functionally different? Novel proteins can evolve
through the shuffling of ancient functional domains into new
combinations (Putnam et al. 2007; Moore et al. 2008). We
therefore asked whether the different proteins found in the
nematocysts from the three cnidarian lineages might be composed of similar domains, potentially leading to a similar functional profile. Indeed, 49 domains are shared between the
three organisms (7.7% of the total number of InterPro
domains identified) including domains involved in protein–
protein, protein–carbohydrate, protein–nucleotide, and protein–membrane interactions (fig. 2B and C). Although relatively few domains are common to all organisms, these
common domains tend to be the most abundant ones detected. The abundance of the domains in the nematocyst
proteome is markedly different from their abundance in the
total predicted Hydra proteome (fig. 2D), further supporting
the hypothesis that these domains represent functions enriched in the nematocysts of all three organisms. Notably,
there are many domains that are not shared by all organisms
but nevertheless belong to the same functional categories—
for example, in addition to the “core” peptidase domains
there are also many organism-specific peptidases. Thus, although the specific proteins found in the nematocysts may
differ between Hydra, Aurelia and Anemonia, the functional
744
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modules from which these proteomes are constructed are
similar.
The Unusual Evolution of Cnidarian Venom Toxins
We next focused on proteins belonging to known toxin families, which are likely major contributors to the “business end”
of the nematocyst, causing prey paralysis or predator deterrence. Given that we are comparing organisms from three
orders that diverged over 500 Ma, how similar is their venom?
Can we identify evolutionary processes that shape the chemical arsenal of these organisms over extended evolutionary
time scales?
As shown in figure 3A, the proteins suggested to comprise
the nematocyst venom arsenal differ between Anemonia,
Aurelia and Hydra, both in terms of the major classes of
venom toxins present in the nematocysts of each animal
(e.g., neurotoxins, cytolysins and enzymes) and in terms of
specific proteins or protein families. Importantly, we observed
that the venoms of the Hydra and Aurelia appear to be functionally similar, whereas the venom of the Anemonia is markedly unique. The Anemonia venom is rich in neurotoxins,
with ten toxins identified belonging to three families of K+
channel toxins and one family of Na+ channel toxins. In contrast, the Aurelia and Hydra venoms contain no identifiable
peptide neurotoxins, but are rich in cytolysins and enzymes.
This broad distinction between Anthozoan and Medosozoan
venoms is supported by several studies using bioassay-guided
fractionation, proteomics, and bioinformatics, which failed to
identify clear orthologs of low molecular weight neurotoxins
in Hydra and various jellyfish or box jellyfish (Klug et al. 1989;
Sher et al. 2005; Brinkman et al. 2012; Li et al. 2012; Badre
2014). Nevertheless, the venoms of Anemonia, Hydra, and
Aurelia are all used to paralyze prey and the stings of
Anemonia and Aurelia are also known to cause pain
(Trembley 1744; Maretić and Russell 1983; Klug et al. 1989;
Chintiroglou and Koukouras 1992; Radwan et al. 2001; Sher
and Zlotkin 2009; Mariottini and Pane 2010). This suggests
that, in each of these organisms, the venom has evolved different pharmacological solutions to the same ecological
needs.
Two potential toxin families that are common to all three
taxa are those containing Membrane Attack Complex/
Perforin (MAC-PF) domains and ShK neurotoxin-like domains. Both of these domains are ancient and widely distributed, with ShK domain found in algae (e.g., UniProt entry
MICPUN_60192 from Micromonas sp) whereas MAC-PF proteins are known to play a role in the pathology of bacteria
(Rosado et al. 2008). MAC-PF domains are pore-forming domains previously identified in hemolytic toxins from the sea
anemone Actineria villosa (Oshiro et al. 2004). The MAC-PF
domains belong to several phylogenetically cohesive groups
that are quite distinct in terms of their sequence, with most of
the anthozan and medusozoan sequences clustering separately and also having different domain structures (fig. 3B,
supplementary figs. S7 and S8, Supplementary Material
online). Some Hydra MAC-PF-containing proteins are also
found in nonnematocystic tissues (Amimoto et al. 2006;
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Nematocyst Content . doi:10.1093/molbev/msu335
A
Orthologs of known cnidarian toxins
K+
Na+
Neurotoxins
Type I Type II Type III
(ShK-like) (Kunitz) (BDS-like)
B
Neurotoxins
Type I
(Av2)
Cytolysins
CaTX-like Actinoporins
*
*
*
ShK-domains MAC-PF Lectin
ATPasewith
like
Number of
Other
(Av120) Metalloproteins
protease
1
4
2
5
3
6
*
*
Complement
*
*
*
*
PLA2
C
*
*
*
*
Putative novel toxins
Perforin
CaTX Kunitz
family
*
*
*
Ancestral gene in
genome/transcriptome
Actinoporin
Toxin-like gene in
genome/transcriptome
Toxin protein in
nematocyst
0.4
FIG. 3. The putative toxic arsenal of Anemonia, Aurelia and Hydra. (A) The presence of orthologs of known cnidarian toxins and of proteins similar to
toxins from other animals is shown superimposed on the phylogeny of the three cnidarians. The size of the circles represents the number of different
proteins identified by MS/MS. The nomenclature of K+ channel toxins follows that of Castaneda and Harvey (2009). (B) Phylogeny of MAC-PF
containing venom proteins from cnidarians venoms and mammalian complement and perforin. A midpoint-rooted ML tree is shown, nodes with
greater than 50% support in both ML and Bayesian analyses are noted by *. Anemonia, Aurelia, and Hydra proteins are marked with green, blue, and
yellow, respectively. The sequence identifiers and multiple alignment used to produce the tree are shown in supplementary figure S7, Supplementary
Material online, and the domain structures are shown in supplementary figure S8, Supplementary Material online. The sequences identified in
Nematostella vectensis have not been identified in the nematocysts (Moran et al. 2013). (C) Potential gene loss and protein derecruitment events
in the evolution of cnidarian venom. A schematic phylogram is shown, with squares denoting 1) the presence in the genome or expression in the
transcriptome of an ancestral gene which is not an RBBH of a toxin (is more similar to endogenous proteins than to toxins) and therefore likely plays an
endogenous role (yellow); 2) a toxin-like gene (orange); 3) the presence of the putative toxin protein in the nematocyst proteome (red).
Miller et al. 2007), and therefore these proteins may have dual
or multiple roles. Assuming the MAC-PF-containing proteins
are indeed toxins, it is currently unclear whether they were
part of the arsenal of the common cnidarian ancestor, or
whether they were recruited independently after the separation of the Anthozoan and Medusozoan lineages, as suggested for toxin families from other venomous lineages (Fry
et al. 2006).
Sea anemones are known to contain type-I ShK-like K+
channel toxins as a part of their venom arsenal (Castaneda
and Harvey 2009). These toxins have a distinct secretion signal
sequence and cleavable pro-region and a single mature sequence. In contrast, the Aurelia and Hydra venoms contain
no identifiable ShK toxins, but include many ShK-like domains in larger proteins, often with other domains such as
Zn metalloproteases. Phylogenetic analysis reveals a marked
distinction between ShK toxins and ShK domains: Although
the Actiniaria toxins are grouped tightly together, the ShK
domains are mostly quite diverse and their phylogeny is
difficult to resolve (supplementary figs. S9 and S10,
Supplementary Material online). Additionally, Rangaraju
et al. (2010) have shown that the ShK domain found as
part of the human matrix metalloprotease 23 can specifically
block K+ channels, but requires concentrations several orders
of magnitude higher than the ShK toxin itself. Thus, it is currently unclear whether the ShK-containing proteins found in
Aurelia and Hydra are in fact toxins, or whether they fulfill
other roles. As toxins, they could directly block K+ channels,
or the ShK domains could serve as “affinity probes” to target
multifunctional proteins to the nerve terminals of the prey or
predators where other domains (e.g., metalloproteases) can
have toxic effects as suggested for the snake venom toxin
-bungarotoxin, (Rowan 2001). Importantly, matrix metalloproteases can be potent toxins in their own right even without ShK domains, and are perhaps one of the bestcharacterized families of snake venom toxins (Fry and
Wuster 2004; Markland and Swenson 2013). Further research
is required to determine to what extent venom activities
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associated with metalloproteases, such as prevention of blood
clotting, inflammation, and apoptosis, are indeed observed in
cnidarian venoms.
Although the MAC-PF and ShK domains are common to
all three organisms, the vast majority of putative toxin families
are found either in the Anemonia or in both Hydra and
Aurelia. Thus, as illustrated most clearly with the low molecular weight neurotoxins, functional recruitment and innovation into the venoms appears to have occurred primarily after
the split between the ancestor of Anemonia and the
common ancestor of Hydra and Aurelia. Furthermore although it is difficult to draw conclusions from only two organisms, it seems that relatively few new toxin families were
recruited into the Aurelia and Hydra venoms after their ancestors had split, even though this occurred not long after the
split between Anthozoans and their common ancestor (fig. 1;
Park et al. 2012). It is tempting to speculate that a significant
evolutionary event occurring during the Ediacaran period,
such as the Avalon Explosion (Shen et al. 2008), had an important part in shaping the venom arsenal of these organisms,
with relatively few recruitment events following this period.
Further research, including targeted studies of the nematocyst
venom of anthozoans other than anemones (e.g., corals and
coralimorpharians), will be required to identify more exactly
when specific toxins families such as the low molecular
weight neurotoxins were recruited into the venom.
How have the cnidarian toxin families evolved following
their recruitment? In many venomous taxa, toxins evolve
through a combination of gene recruitment, duplication,
and neofunctionalization, resulting in extended families of
closely related yet often functionally distinct toxin isoforms
which reveal molecular hallmarks of positive selection (Duda
and Palumbi 1999; Fry et al. 2009; Kozminsky-Atias and
Zilberberg 2012; Casewell et al. 2013). Although for some
families moderate gene multiplication could be observed,
none of the toxin families we identified exhibited clear hallmarks of gene diversification through positive selection, or
any signal of diversifying selection postrecruitment (supplementary figs. S11–S14 and Excel file S2, Supplementary
Material online). This is puzzling, since according to VanValen’s Red Queen Hypothesis (Van Valen 1974), to remain
effective, toxins used for predation are required to evolve
constantly to counteract the acquisition of resistance by
prey (“molecular arms race,” e.g., Poran et al. 1987).
However, this may be a result of several factors. First, the
sample size for analysis is somewhat limited. In most analyses
alignments contained approximately ten sequences, mostly
belonging to the same species, which may skew the analysis,
compared with a broader sampling of various families.
Second, not all toxins are expected to be under positive selection. Indeed Kozminsky-Atias and Zilberberg (2012) have
shown that the toxin population within a species shows varying signals of pressure of selection. Finally, although positive
selection has often been demonstrated for neurotoxins,
toxins targeting lipids and other molecules whose production
machinery is complex show considerably lesser footprint of
positive selection. For example, a thorough and comprehensive analysis of spider sphingomyelinase D by Binford et al.
746
(2009) found marginal evidence for positive selection, with
high purifying selection acting on the active sites of these
enzymes. Thus, in the case of the cholesterol targeting
MAC-PF and the phospholipid targeting pore-forming
toxins, an overall purifying selection should be expected.
Still, it is surprising to find insignificant level of positive selection in Actiniaria type 2 and 3 K+ channel neurotoxins, and
this may suggest that cnidarian venoms are not exposed to
the selection pressure associated with a molecular arms race,
or that the cnidarian’s venom arsenal evolves in a different
manner, that does not leave the molecular traces of positive
selection. In the case of ShK-containing proteins, in which the
ShK domain (in Hydra) is encoded on a separate exon
(Rachamim and Sher 2012), new functions may be imparted
through new domain combinations, a common mechanism
suggested to occur in many eukaryotic proteins such as receptors and signaling ligands (Moore et al. 2008; supplementary fig. S10, Supplementary Material online).
Finally, in parallel to the emergence of new toxin families,
our data suggest that several toxin families seem to have been
lost from the venom of specific cnidarian lineages (fig. 3A
and C). For example, proteins similar to the CaTX family of
hemolysins were found in the venom of Hydra and Aurelia,
defining a novel clade of CaTX-like toxins, but could not be
identified in Anemonia. Nevertheless, a gene encoding a
CaTX-like transcript was identified in the Anemonia transcriptome (supplementary figs. S15 and S16, Supplementary
Material online). Similarly, hemolytic actinoporin-like proteins
were identified in the nematocysts of Hydra (Glasser et al.
2014) but were not identified in the nematocysts of Aurelia
and Anemonia, despite this toxin family being common (and
intensively studied) in sea anemones from the anemone
family to which Anemonia belongs (Actiniidae, (Kristan
et al. 2009). We could not identify a homologous gene in
the transcriptome of Aurelia, or in the recently published
transcriptome from a different Aurelia strain (Fuchs et al.
2014). In contrast, the transcriptome of Anemonia contains
a long protein with two actinoporin-like domains in tandem
on the same transcript. Given that the N0 -terminus of actinoporins is the part that inserts into the membrane to line a
transmembrane pore (Athanasiadis et al. 2001; Manche~no
et al. 2003), it is likely that the fusion protein has lost some
or even all of its hemolytic activity. Taken together, these
results suggest that selective loss or “decommissioning” of
protein families from the venom and potentially from the
genome itself is common, with actinoporins likely being lost
from the lineage of Anemonia relatively recently.
The Dynamic Nematocyst
Nematocysts are ancient organelles, which perform a complex function of explosive discharge, one of the fastest cellular
events in nature. One might expect that multiple proteins
would need to work closely together to produce such an
effect and indeed the basic structure and activation mechanism are relatively conserved among nematocysts from various species (Tardent 1995). However, our findings show that
this functional similarity is not mirrored by similarity in the
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soluble protein payload between nematocysts from different
organisms. Rather, the nematocyst payload is unique to each
organism and potentially to each nematocyst type (Hwang
et al. 2007). It is possible that the insoluble fraction of the
nematocyst proteome, which likely comprises the majority of
the structural proteins in the nematocyst, is more similar
between the three organisms. Additionally, the diversity of
morphological forms and life histories among cnidarians is
immense, and clearly the three organisms studied here
cannot be taken as representatives of the entire phylum.
Nevertheless, for the nematocysts from these three model
organisms, we propose that, over long time scales of hundreds
of millions of years, different combinations of similar modules
(protein domains) have likely evolved to perform similar or
related functions. Some of the proteins in the payload are
likely involved in nematocyst assembly, and are now trapped
in the mature organelle; others, for example GGH, may be
involved in nematocyst discharge, whereas a relatively small
percent (3–7% of the total nematocyst proteome) are likely
“effector proteins,” namely toxins. Much of the toxin arsenal
seems to have evolved after the separation of the three cnidarian lineages. Therefore, although the injection of cnidarian
venom typically causes almost immediate prey paralysis,
with the possible exception of the effects of similar MACPF-containing proteins, the venom of each of the cnidarian
lineages likely causes this effect mostly through distinct pharmacological mechanisms. Notably, additional toxins, not
found in the nematocysts themselves, may also take part in
prey capture or predator deterrence, with their exact delivery
mechanism still unclear (Moran et al. 2011). Why the evolutionary patterns of cnidarians venom differ from many other
venomous organisms is still unclear. We speculate that the
intricate manner in which venom production is presumably
tied in and coordinated with the nematocyst development, to
allow the toxins to be packed in the developing nematocyst,
might pose unique developmental or genomic constraints on
venom evolution in cnidarians compared with other venomous organisms. Such constraints may lead to an abundance of
bifunctional proteins with roles in both nematocyst development and toxicity, as well as restrict the emergence of a molecular “arms race” between the cnidarians and its prey.
Without the ability to win an “evolutionary arms race,” a
significant portion of the cnidarians venoms may have
evolved to hijack endogenous mechanisms such as activation
of the endothelin system, or alternatively target the cell membrane or extracellular matrix (e.g., through pore-forming
toxins, phospholipases, and metalloproteases), against
which the development of resistance might be significantly
more difficult.
Materials and Methods
Organisms
Hydra magnipapillata (a kind gift of Prof. Thomas Bosch,
University of Kiel) was cultured at 18 C in buffer M (1 mM
CaCl2, 1 mM Tris–HCl pH 7.7, 0.1 mM KCl, 0.1 mM MgCl2,
1 mM NaHCO3) and fed every other day with freshly hatched
Artemia nauplii. Aurelia aurita was collected from a bloom in
the gulf of Eilat during February–April 2012. The organisms
were immediately dissected to separate the tentacles from
the bell, and were frozen at 80 C until nematocysts were
isolated. Anemonia viridis was collected at Sdot Yam (Israel),
brought to the lab, and maintained in aquaria at room temperature for several months.
Isolation of Hydra Nematocysts
The isolation of the nematocyst from all organisms was performed by modifications of a similar protocol based on lysis of
the tissue followed by isolating the dense nematocysts by
centrifugation on a bed of Percoll, which differentiates between the dense intact nematocyst microcapsules and the
cell debris (Marchini et al. 2004; Ayalon et al. 2011; Tal et al.
2014). Hydra were starved for at least 72 h. Animals were
gently picked with Pasteur pipette into cold Distilled Water
(DW), and placed on ice for 30 min. The hydra than transferred to a clean 15-ml tube with fresh cold DW, allowed to
settle, and the pellet frozen at 20 C and then transferred to
80 C until used. To isolate the nematocysts, frozen hydra
were thawed in 5 ml of 10% glycerol, 50% Percoll, followed by
short vortex, and pipetting. The tube was placed on ice for
30 min, centrifuged at 6 C, 3,000 g for 15 min, the upper
fraction was discarded, and the lower fraction was resuspended in the same buffer with the addition of 0.005%
Triton x-100. This was repeated once in the same buffer
and twice more in DW, with the final and pure nematocyst
pellet resuspended in 50 ml Double Distilled Water (DDW).
Isolation of Anemonia Nematocysts
Tentacles were cut on ice, homogenized with sea water and
the homogenate was shaken at room temperature for 1 h;
500 ml of the homogenate was placed on top of 1 ml 70%
Percoll, followed by centrifugation at 6 C 3,000 g for
10 min. The pellet was suspended in fresh sea water with
7% sucrose, placed on top of 1 ml of 10% glycerol, 85%
Percoll, and centrifuged at 6 C, 3,200 g for 70 min. The
pellet was resuspended in 1 ml of 5% Triton x-100 in sea
water, followed by centrifugation and washing twice in DW
as described above for the nematocysts from Hydra.
Isolation of Aurelia Nematocysts
The frozen tentacles were defrosted and homogenized in 50%
Percoll and kept on ice for 25 min. Tritonx-100 was added to a
final concentration of 0.1%, the sample left on ice for 5 min
and centrifuged at room temperature, 800 g for 10 min. The
resulting pellet was resuspended in 2 ml of 0.1% Tritonx-100,
and 48 ml of DDW, centrifuged (800 g, room temperature
(RT), 10 min), and the process repeated once in the same
buffer, four more times in 0.5 ml of 0.1% Triton x-100 (with
the last three centrifugations lasting 5 min) and twice more in
DDW. The final pellet was resuspended in 200 ml of DDW.
Nucleic Acid Isolation, Sequencing, and Assembly
RNA was isolated from several different developmental stages
of Aurelia (planula, polyp, early and late strobili, ephyra, and
the gonads of mature jellyfish, [Brekhman et al. 2014]) and
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Anemonia (tentacles, pharynx, septae, and body wall) using
TriReagent (Sigma), and Illumina libraries were prepared
using the TruSeq kits (Illumina). Sequencing (100 bp, paired
end) was performed on a HiSeq2000 sequencer (Illumina) at
the LS&E unit of the Technion, Israel. A full description of the
assembly and annotation of the Aurelia transcriptome is presented elsewhere (Brekhman et al. 2014). The transcriptome
of Anemonia was assembled using Trinity (Grabherr et al.
2011) with the default parameters (K-mer = 25). We also sequenced and partially assembled, using the same sequencing
technology, the genome of Anemonia, with the aim of testing
whether the low-coverage, fragmented genomes produced
using our approach would increase the number of proteins
identified by shotgun MS/MS. This assembly was performed
using SOAPdenovo (Li et al. 2009) with a K-mer of 31 for
assembly and 25 for GapCloser. The final reference databases
used for interpreting the MS/MS spectra were as follows:
H. magnipapillata: 17,188 translated proteins from the published genome sequence (Chapman et al. 2010); Au. aurita:
Approximately 322 Mb of transcriptomic sequences; A. viridis:
Approximately 230 Mb and approximately 75 Mb of genomic
and transcriptomic sequences, together with approximately
40,000 published expressed sequence tag (ESTs) (see below).
The sequencing reads and assembled transcriptome of
Anemonia were deposited in GenBank as bioprojects
PRJNA260824 (Anemonia).
Selecting Nucleic Acid Databases for MS/MS Analysis
Anemonia has been studied extensively as a model for cnidarians-dinoflagellate symbiosis, and approximately 40,000
ESTs are available (Sabourault et al. 2009). This enabled us
to test the quality of our rough transcriptome and genome
assemblies, and to determine which molecule—genomic
DNA or mRNA—is more appropriate for sequencing with
the aim of downstream proteomic analyses. As shown in
supplementary figure S1, Supplementary Material online,
almost all of the published Anemonia ESTs were covered to
some extent by both the transcriptome and genome assemblies (99.95% and 91% for the transcriptome and genome,
respectively). The transcriptome assembly outperformed the
genome assembly both in terms of covering the known ESTs
(supplementary fig. S1A and B, Supplementary Material
online) and in terms of enabling the identification of more
proteins using MS/MS (supplementary fig. S1C, Supplementary Material online). The better performance of the transcriptome and genome assemblies compared with the
published ESTs in identifying proteins using MS/MS may be
due in part to better sequence coverage and in part to using
RNA and DNA from the same specimen or population used
for the MS/MS. Nevertheless, the combination of transcriptome, genome, and EST databases enabled the identification
of more proteins than either of these databases alone
(supplementary fig. S1, Supplementary Material online). As
the genomic and transcriptomic databases were to be used
specifically for subsequent MS/MS analysis, we did not perform further systemic postassembly analyses such as
annotation.
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Mass Spectrometry
Prior to MS/MS, the isolated nematocysts were physically
ruptured using a TissueLyser II bead beater (Qiagen) using
1-mm glass beads at maximum shaking speed for 10 min,
using no chaotropic agents. MS/MS was performed at the
Smoler Center for Peropteomics, Technion, Israel. The soluble
nematocyst content in 8 M urea and 100 mM ammonium
bicarbonate was reduced with 2.8 mM DTT (60 C for
30 min), alkylated in the dark at room temperature with
8.8 mM iodoacetamide in 100 mM ammonium bicarbonate,
and digested overnight at 37 C in 2 M urea, 25 mM ammonium bicarbonate with TCPK-treated trypsin (Promega) at a
1:50 enzyme-to-substrate ratio. The tryptic peptides were desalted using homemade C18 stage tips, dried and re-suspended in 0.1% Formic acid, and resolved by reverse-phase
chromatography on 0.075 200 mM fused silica capillaries
(J&W) packed with Reprosil reversed phase C18 material
(Dr Maisch GmbH, Germany). The peptides were eluted
with linear 95 min gradient of 7–40% and 8 min to 95% acetonitrile with 0.1% formic acid in water at flow rates of 0.25
ml/min. MS/MS was performed by a hybrid LTQ-Orbitrap XL
(Orbitrap, Thermo) in a positive mode. High resolution MS1
scans in the orbitrap were performed at 60K resolution, followed by collision-induced dissociation of the seven most
abundant ions and low resolution scans in the ion trap.
Data were searched using Byonic (Protein Metrics) software
against a six-frame translation of the Aurelia and Anemonia
combined databases, and against the Hydra Open Reading
Frames (ORFs) derived from the genome. The search was
performed using semispecific enzyme cleavage with two
missed cleavages, with 10 ppm mass tolerance for the precursor and 0.7 Da for the fragment ions, with 100 Da wildcard
(mass deviation from reference sequence). Search was run
against a decoy database with tolerance of 1% false discovery
rate. Hits with less than two unique peptides per protein were
dismissed. The identified proteins (accession numbers, sequences, Gene Ontology (GO) annotations, and InterPro results), as well as the results of the Byonic searches, are found in
the supplementary Excel file, Supplementary Material online.
Assessing the Purity of the Nematocyst Proteomes
The nematocysts were pure from contaminating cellular
debris by light microscopy (Fig 1), with the exception of the
Anemonia nematocysts, which coeluted from the density
centrifugation with a small amount of symbiotic algae
which have a similar size and density. To provide further
evidence of the purity of the nematocysts from Hydra, we
used BLASTp to search the nematocyst proteome for 25 proteins known to be expressed exclusively in nonnematocystic
tissues (gland cells, i-cells, and neurons, taken from supplementary tables S4–S6 in Hwang et al. [2007]). None of these
25 proteins was identified in the nematocyst proteome (defined as 4 95% sequence identity), suggesting that the nematocysts are essentially free of significant cellular
contaminants. To estimate the level of contamination of
the Anemonia nematocyst proteome by proteins from the
symbiotic algae, we used tBLASTn to search the genomes of
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Symbiodinium minutum Mf 1.05b.01 (a culturable zooxanthellae originally isolated from the coral Montastraea faveolata; Shoguchi et al. 2013) and the nonsymbiotic sea anemone
Nematostella vectensis (Putnam et al. 2007) for the proteins
identified in the Anemonia nematocysts. Of the 374 proteins
identified in the nematocysts, 93 had BLAST hits to
Symbiodinium and 203 to Nematostella. Of these, 11 were
identified in the Symbiodinium genome but not in the
Nematostella genome, and an additional 26 had a higher
bit-score in the tBLASTn search against the Symbiodinium
compared with the Nematostella genome. Thus, 37/374 proteins (or approximately 10%) are likely contaminants from
the algae. Most of these proteins are involved in photosynthesis or are general, high-abundance housekeeping genes
such as ribosomal proteins, myosin or the enzyme glyceraldehyde-3-phosphate dehydrogenase. Given that the origin of
the proteins is still unclear, that they comprise only approximately 10% of the proteome, that none of them was an
RBBH and that similar proteins were usually found that
were not suspected to be contaminants, we retained these
proteins in downstream analyses but highlight them in the
supplementary Excel file S1, Supplementary Material online.
Comparison of Our Results with a Previously Released
Nematocyst Proteome from Hydra
We tested how well our shotgun MS/MS approach performs
compared with a recent in-depth analysis of the Hydra nematocyst content (Balasubramanian et al. 2012). Direct comparison of the results obtained with both methods is difficult,
as, in addition to differences in the starting material and proteomic methods, the Hydra gene models used for identifying
the protein spectra were obtained from different sources (the
Hydra genome browser Hma2 models for Chapman et al.
[2010]; Balasubramanian et al. [2012]) and the Hydra protein
from the NCBI nr database for our analysis) and were not
computationally connected. Nevertheless, depending on the
MS/MS algorithm used and the definition of “identical” proteins (what percent coverage and what percent identify), we
detected approximately 33–69% of the proteins identified by
Balasubramanian et al. (2012) using significantly fewer massspectrometry analyses (supplementary fig. S2, Supplementary
Material online).
Comparing Proteomes
The sequences identified as part of the nematocyst payload
by the MS/MS analysis were extracted from the full transcriptomes, translated when necessary into protein sequences, and
compared with identify RBBHs (Altenhoff and Dessimoz
2009) using in-house BLASTp (protein vs. protein database)
searches. Ten putative RBBHs were identified, and this
number was used to calculate the abundance of RBBHs in
the nematocysts compared with the full Hydra proteome.
These ten sequences were then manually inspected to identify whether the region of similarity extended over at most of
the protein (defined as a BLAST hit over at least 60% of the
coding sequence of the longest coding sequence), resulting in
the six full-protein RBBHs shown in table 1. A similar analysis
of the entire Hydra proteome was performed by searching for
homologs of 17,580 putative Hydra proteins (downloaded
from the NCBI nr database during January 2013) in the
Anemonia and Aurelia transcriptomes using tBLASTn (protein vs. six-frame translated database), followed by reciprocal
BLASTx (nucleotide vs. protein database). The subset of sequences that were RBBHs between Hydra and both
Anemonia and Aurelia was then tested by tBLASTx (translated nucleotide vs. translated nucleotide) to determine
whether they are also RBBHs between Anemonia and
Aurelia. Out of 17,580 hydra proteins, 4,690 are RBBHs between Hydra, Anemonia, and Aurelia. To identify common
functionalities, BLAST2GO (Conesa et al. 2005) was used to
obtain the InterPro domains found in the various nematocyst
proteins (October 2014 release), with comparisons performed
using Microsoft Excel.
Identifying Toxins in Nematocyst Proteomes
The proteins identified by the MS/MS analysis were analyzed
using a pipeline aimed at conservatively identifying toxins:
Proteins which are true orthologs (RBBHs; Altenhoff and
Dessimoz 2009) of cnidarian toxins were deemed more
likely to be bona fide toxins than those that are similar to
toxins from distantly related organisms. We first searched the
nematocyst proteomes using, as the BLASTp query, the nonredundant list of 191 Anemone toxins from Oliveira et al.
(2012), with the addition of the following toxin sequences
from Hydrozoans, Schyphozoans, and Box Jellyfish: Hln1 and
Hln3 (H. viridissima, accession numbers Q86LR2.3, Q52SK6.3),
Toxin A (Carybdeaalata, Q9GNN8.1), CqTX-A (Chiropsalmus
quadrigatus, P58762.1), CfTx-1 and 2 (C. fleckeri, A7L035.1,
A7L036.1), CrTX-A (Carybdearastonii, Q9GV72.1), Millepora
cytotoxin-1 (Millepora dichotoma, A8QZJ5.1), and the partial
sequence of a PLA2 from Rhopilema nomadica (P43318.1).
BLASTs were performed in-house using the command-line
BLAST version 2.2.29, the default BLOSUM62 matrix, and
an e value cutoff of 10e5. To verify that the toxin-like sequences in the nematocysts are indeed more similar to cnidarians toxins than to endogenous toxin-like proteins, we
determined whether the sequences are RBBHs against the
SwissProt protein database at the NCBI (September 2014 release), and only these sequences were annotated as cnidarians
toxins. Subsequently, a similar RBBH analysis was performed
using, as query, all toxins downloaded on September 24, 2014
from the ToxProt database. Two homologs of Avt-120 (Uechi
et al. 2011) from Aurelia were identified as toxins based on the
BLAST2GO annotation, and added to the list of known cnidarians toxins. Finally, we added to the list of putative toxins
proteins with metalloprotease, ShK or MAC-PF domains, as
these have previously been suggested to form part of the
cnidarian chemical arsenal (Oshiro et al. 2004; Moran et al.
2013; Rachamim and Sher 2012). We chose not to include in
our putative toxin list candidates proteins with domains commonly associated with toxins such as CRISP or CAP domains
(Fry et al. 2009), as our data set contained many such proteins,
for which there is no experimental evidence of toxicity.
Furthermore, preliminary searches showed that the
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nematocyst proteins containing CRISP domains are all relatively large, lack the ICR (Ion Channel Regulatory) domain
which binds ion channels, and are not RBBHs of venom proteins. Thus, although the toxic activity of the proteins containing CRISP domains cannot be precluded we did not
analyze them further in this study.
Marine Sciences to T.R. Sequencing was partly funded by
the RBNI program at the Technion, Israel. The authors declare
no competing interests. T.R., T.L., and D.S. designed research;
T.R., D.M., D.A., and V.B. performed research; T.R., D.M., T.L.,
and D.S. analyzed data and wrote the paper.
References
Phylogenetic and Selection Analyses
Protein sequences were aligned using either ClustalW,
MUSCLE or MAFFT, the latter using E-INS-i or L-INS-I algorithms, and corrected manually (in MEGA 5.0; Tamura et al.
2007). Codon alignment was then generated by superimposing the nucleic acid data over the protein alignment using
DAMBE (Xia 2013), followed by saturation analysis of the
alignment using the same software. We generally performed
phylogenetic analysis using both Maximum Likelihood (ML)
and Bayesian methods. For ML analysis, amino acid substitution model was assessed using ProtTest 3.0 (Darriba et al.
2011) and the trees generated in MEGA 5.0 with 1,000 bootstrap pseudoreplications. The selected models were
WAG+I+G+F for the ECEs and CaTX-like proteins, WAG+F
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Supplementary Material
Supplementary Excel files S1 and S2 and figures S1–S16 are
available at Molecular Biology and Evolution online (http://
www.mbe.oxfordjournals.org/).
Acknowledgments
The authors thank Hila Wolf (Technion, Israel) for performing
mass spectrometry analysis, Brian J. Haas (Broad Institute,
USA), Leonid Brodsky and Amir Cohanim (University of
Haifa, Israel) for help with transcriptome assembly, Assaf
Malik and Noa Sher (University of Haifa Bioinformatics
Support Unit) for help with bioinformatics analyses, and Uri
Shavit (Technion, Israel) for assistance and collaboration.
They also thank three anonymous referees who contributed
significantly to this manuscript. This work was supported by
grant 1994/13 from the Israel Science Foundation to D.S. and
by a postdoctoral fellowship from the Charney School of
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