Aquatic Toxicology 104 (2011) 61–72
Contents lists available at ScienceDirect
Aquatic Toxicology
journal homepage: www.elsevier.com/locate/aquatox
Review
On the origins and biosynthesis of tetrodotoxin
Rocky Chau, John A. Kalaitzis, Brett A. Neilan ∗
School of Biotechnology and Biomolecular Sciences, The University of New South Wales, Sydney, NSW 2052, Australia
a r t i c l e
i n f o
Article history:
Received 18 November 2010
Received in revised form 30 March 2011
Accepted 1 April 2011
Keywords:
Tetrodotoxin (TTX)
Toxin biosynthesis
Polyketide
Non-ribosomal peptide
Amidinotransferase
a b s t r a c t
The potent neurotoxin tetrodotoxin (TTX) has been identified from taxonomically diverse marine organisms. TTX possesses a unique cage-like structure, however, its biosynthesis has yet to be elucidated.
Biosynthetic studies in the TTX-producing newt Taricha torosa, and in bacterial genera, including Vibrio, have proven inconclusive. Indeed, very few studies have been performed that address the cellular
production of TTX. Here we review the sources of TTX described to date and provide evidence for the
biosynthesis of TTX by symbiotic microorganisms in higher taxa. Chemical and genetic based biosynthesis studies of TTX undertaken thus far are discussed and we outline approaches which may be useful for
expanding upon the current body of knowledge. The complex biosynthesis of structurally similar toxins,
that reveal clues into the biosynthetic pathway of TTX, is also presented.
© 2011 Elsevier B.V. All rights reserved.
Contents
1.
2.
3.
4.
5.
Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Tetrodotoxin overview . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
2.1.
Toxicity of tetrodotoxin . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
2.2.
Structure of tetrodotoxin . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
2.3.
Supply of tetrodotoxin . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Distribution and biogenic origins of tetrodotoxin . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
3.1.
Bacterial origins of tetrodotoxin . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
3.2.
Non-bacterial origins of tetrodotoxin . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
3.3.
Dietary origins of tetrodotoxin in higher animals . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Biosynthesis of tetrodotoxin . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
4.1.
Feeding experiments . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
4.2.
The genetic basis for a proposed tetrodotoxin biosynthesis pathway . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
4.2.1.
The guanidinium moiety of tetrodotoxin . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
4.2.2.
The carbon backbone of tetrodotoxin . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Future directions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
1. Introduction
The oceans are a rich source of biologically active and structurally novel natural products derived from a myriad of organisms
and biosynthetic pathways (Blunt et al., 2010). Many of these
natural products serve important ecological roles in the marine
environment, such as species survival, and act as chemical defenses,
camouflage agents, or anti-foulants (Hay and Fenical, 1996). The
bioactivities of such compounds have been adapted for use by
∗ Corresponding author.
E-mail address: b.neilan@unsw.edu.au (B.A. Neilan).
0166-445X/$ – see front matter © 2011 Elsevier B.V. All rights reserved.
doi:10.1016/j.aquatox.2011.04.001
61
62
62
63
63
64
64
65
66
66
66
66
67
69
69
70
the pharmaceutical and biotech industries to drive drug discovery and develop chemical tools for the benefit of humans (Glaser
and Mayer, 2009). In contrast, the same complement of organisms biosynthesize some of the most toxic substances known to
mankind, ranging from those with neurotoxic and paralytic effects
to others causing skin irritations (Kalaitzis et al., 2010). Many
marine toxins possess highly unusual chemical structures (Fig. 1),
resulting from complex biosynthetic pathways active in the producing organism. In some cases even proposing a biosynthetic
route to the toxin has proved difficult due to the structural complexity of the compound (Kalaitzis et al., 2010). Traditionally, such
pathways were proposed based on feeding experiments whereby
radioactive isotope labelled precursors were fed to the producing
62
R. Chau et al. / Aquatic Toxicology 104 (2011) 61–72
Fig. 1. Structures of natural products discussed in this review.
organism (Minor et al., 1954). Detection of radioactivity associated with the purified target molecule confirmed incorporation
of the labelled precursor. Improvements in NMR technologies led
to it being used as the method of choice for monitoring stable
isotope incorporation. Its use not only allowed for levels of incorporation to be quantified but it also had the added advantage of
locating the labels on the molecule through comparison with the
unlabelled substance (Bacher et al., 1998). These analyses revealed
the fate of the precursor through the biosynthetic pathway to
the final product and locations of the labels from various substrates helped to ultimately determine the proposed biosynthesis
pathway.
Biosynthesis studies have undergone a re-birthing of sorts with
the development of modern day molecular genetics techniques to
aid the elucidation of toxin biosynthesis pathways. A lot is now
known about the genetics underpinning natural product assembly, particularly in microorganisms (Cane and Walsh, 1999). Genes
coding for natural product biosynthesis enzymes, as well as those
associated with regulation and transport of the natural product,
particularly toxins, are generally clustered on bacterial genomes.
As a direct result of this, biosynthesis gene clusters are highly
amenable to detection using molecular tools (Udwary et al., 2007).
Identification and complete characterization of these biosynthetic
gene clusters has, in recent times, become a major focus of research
for those investigating natural product biosynthesis pathways. The
elucidation of biosynthesis pathways opens the door to novel
biosynthesis enzymes and the unusual chemistry they catalyze.
One molecule whose assembly remains unproven is tetrodotoxin
(Brown and Mosher, 1963) (TTX—Fig. 2). Tetrodotoxin, whose name
is derived from Tetraodontidae, the family of puffer fish from which
it was first isolated, is also found from other sources including but
not limited to, frogs, the blue-ringed octopus, and molluscs. TTX
is a neurotoxin with a highly unusual structure, whose biosynthesis has yet to be elucidated nearly a century after its discovery.
Such is the cryptic nature of its assembly, elucidation of its path-
way would likely reveal novel biosynthetic reactions and catalytic
enzymes. Furthermore, harnessing the TTX biosynthesis gene cluster by way of cloning and expression would provide a constant
source for commercial use, thus overcoming difficulties associated
with total synthesis. In general, complete characterization and an
intimate knowledge of the biosynthesis gene cluster would also
shed further light on the assembly of similar complex structures in
microorganisms.
Here we present an overview of the approaches employed thus
far to elucidate the TTX biosynthesis pathway. We also discuss
the biosynthesis of structurally related toxins for comparative purposes and highlight the similar pathways that may be in operation
in TTX-producing organisms.
2. Tetrodotoxin overview
2.1. Toxicity of tetrodotoxin
The toxic effects of TTX have been known since antiquity, it is out
of the scope of this review to discuss them at length, however, it is
important to highlight some aspects of its toxicity. Recent reviews
on this topic (Fuhrman, 1986; Narahashi, 2001, 2008) should be
consulted for a complete overview. Historically, both the ancient
Egyptian and Chinese societies possessed knowledge of the toxic
properties of puffer-fish. This is evidenced by engravings of pufferfish found on a 5th dynasty (2500 B.C.) Egyptian tomb (Halstead,
1958; Mills and Passmore, 1988), and reference to the toxicity of
puffer-fish eggs has been recorded as early as the first or second
century B.C. by the Chinese. A more recent (around the year 1600)
description of toxic puffer fish was also provided in the Chinese
materia medica (Fuhrman, 1986). The best known description, by
Europeans, of the toxicity of puffer toxin can be found in the journals of Cook (1775). Hence, it can be seen that the toxic effects of
tetrodotoxin have been well documented throughout history.
R. Chau et al. / Aquatic Toxicology 104 (2011) 61–72
63
Fig. 2. Tetrodotoxin and analogues.
2.2. Structure of tetrodotoxin
Despite knowledge of its toxicity, TTX was not isolated, as a
crystal, until the early 1950s (Yokoo, 1950), and also chromatographically in the 1960s (Brown and Mosher, 1963; Fuhrman,
1986). The structure was not confirmed until several groups elucidated it independent of each other in the mid 1960s (Goto
et al., 1965; Tsuda et al., 1964; Woodward and Gougoutas, 1964).
Although toxicity and pharmacology of TTX are well understood, much of the research regarding the biogenesis of TTX is
still speculative. TTX has a highly unusual structure containing
a single guanidinium moiety attached to a highly oxygenated
carbon backbone. The carbon backbone of TTX consists of a 2,4dioxaadamantane structure, decorated with 5 hydroxyl groups. The
structure of TTX will serve as a basis for the later discussion of its
biosynthesis. TTX and analogues are displayed in Fig. 2.
2.3. Supply of tetrodotoxin
TTX selectively blocks voltage-gated sodium channels, and
because of this selectivity TTX has come to be an important chemical tool in neuroscience (Akopian et al., 1999; Sanford et al., 2006).
Additionally, its use in anaesthesia and analgesia is also being developed (Epstein-Barash et al., 2009; Hagen et al., 2008, 2007; Kohane
et al., 2003) thus heightening its demand. This demand for TTX has
put considerable strain on the current method of TTX isolation and
purification, which involves harvesting of puffer fish livers (Zhou
and Shum, 2003). This isolation method has a detrimental impact
on aquatic life, and is quite inefficient hence other sources of TTX
are required.
Attempts at chemical synthesis of TTX have been successful, but
these generally involve complex, multi-step reactions and result
in low yields. The first successful synthesis was by Kishi et al.
(1972a,b), who reported two different synthetic routes to TTX. Subsequent syntheses have been reported by various research groups
(Table 1), but discussion of these synthetic studies are out of the
scope of this review. As these syntheses in general are time consuming, expensive, and have low yields, a more reliable, cost effective
and sustainable method of TTX supply is sought. This being the
case, harnessing TTX biosynthesis genes in order to produce a
constant and reproducible source of the molecule could be the
most reliable method toward stockpiling TTX. Modern molecular genetic methods have enabled a greater understanding of the
assembly of natural products such as TTX, and a thorough knowledge of it’s biosynthesis at a genetic level will likely allow for
the precise genetic engineering of it in microbes, facilitating the
production TTX in larger quantities that are presently available.
To enable these types of experiments to proceed, it is important
to fully understand the origins and distribution of the molecule
itself.
Table 1
Reported syntheses of TTX.
Authors
Yeara
Synthetic steps
Overall yield (%)
Reference
Y. Kishi et al.
M. Isobe et al.
J. Du Bois et al.
K. Sato et al.
1972
2003
2003
2008
23/26
67
34
34
1.82/1.07
0.64
0.97
0.34
Kishi et al. (1972a, 1972b)
Ohyabu et al. (2003)
Hinman and Du Bois (2003)
Sato et al. (2008)
a
Year reported.
64
R. Chau et al. / Aquatic Toxicology 104 (2011) 61–72
Table 2
Phylogenetic distribution of organisms reported to be sources of TTX (excluding bacteria).
Phylum
Class
Order
Family
Binomial name
Reference
Chordata (includes
frogs, puffer fish,
gobies and newts)
Amphibia
Anura
Dendrobatidae
Brachycephalidae
Colostethus inguinalis
Brachycephalus pernix
Brachycephalus ephippium
Atelopus oxyrhynchus
Notophthalmus viridescens
Taricha torosa
Takifugu xanthopterus
Takifugu niphobles
Fugu rubripesa
Fugu vermicularisa
Fugu pardalisa
Fugu poecilonotusa
Gobius criniger
Hapalochlaena maculosaa
Niotha clathrata
Nassarius semiplicatusa
Rapana rapiformis
Rapana venosa venosa
Charonia sauliae
Babylonia japonica
Polinices didyma
Tutufa lissostoma
Cephalothrix rufifronsa
Lineus longissimusa
Astropecten latespinosus
Astropecten polyacanthusa
Flaccisagitta enflata
Parasagitta elegans
Zonosagitta nagae
Eukrohnia hamata
Carcinoscorpius rotundicauda
Lophozozymus pictor
Atergatis floridus
Planocera multitentaculata
Alexandrium tamarense
Daly et al. (1994)
Pires et al. (2005)
Pires et al. (2002)
Mebs and Schmidt (1989)
Yotsu-Yamashita and Mebs (2003)
Brown and Mosher (1963)
Nagashima et al. (2001)
Yu et al. (2004)
Wu et al. (2005a,b)
Lee et al. (2000), Noguchi et al. (1987)
Yasumoto et al. (1986)
Yasumoto et al. (1986), Yotsu et al. (1987)
Noguchi and Hashimoto (1973)
Hwang et al. (1989), Sheumack et al. (1984)
Jeon et al. (1984)
Wang et al. (2008)
Hwang et al. (1991)
Hwang et al. (1991)
Narita et al. (1981)
Noguchi et al. (1981)
Shiu et al. (2003)
Noguchi et al. (1984)
Carroll et al. (2003)
Carroll et al. (2003)
Maruyama et al. (1984)
Miyazawa et al. (1985)
Thuesen and Kogure (1989)
Thuesen and Kogure (1989)
Thuesen and Kogure (1989)
Thuesen and Kogure (1989)
Dao et al. (2009), Kungsuwan et al. (1987)
Tsai et al. (1995)
Noguchi et al. (1983)
Miyazawa et al. (1986)
Kodama et al. (1996)
Chordata
Mollusca (gastropods)
Bufonidae
Salamandridae
Actinopterygii
Tetraodontiformes Tetraodontidae
Cephalopoda
Gastropoda
Perciformes
Octopoda
Neogastropoda
Gobiidae
Octopodidae
Nassariidae
Muricidae
Stelleroidea
Paleonemertea
Heteronemertea
Paxillosida
Ranellidae
Buccinidae
Naticidae
Bursidae
Cephalothricidae
Lineidae
Astropectinidae
Sagittoidea
Aphragmophora
Sagittidae
Arthropoda (crabs)
Merostomata
Malacostraca
Phragmophora
Xiphosura
Decapoda
Platyhelminthes (flatworms)
Dinoflagellata
Turbellaria
Dinophyceae
Polycladida
Gonyaulacales
Sorbeoconcha
Neotaeniogloss
Nemertea (nematodes)
Anopla
Echinodermata
(starfish)
Chaetognatha (arrow
worms)
a
Pterosagittidae
Eukrohniidae
Cleroidea
Carpiliidae
Xanthoidea
Planoceridae
Goniodomataceae
Bacterial production of TTX reported (Table 3).
3. Distribution and biogenic origins of tetrodotoxin
Over the last 50 years, numerous reports of TTX occurrences
have been published (Table 2). TTX has been found to occur in at
least 6 phyla of organisms within the Animalia kingdom including
the Chordata, Mollusca, Echinodermata, Chaetognatha, Arthropoda
and Platyhelminthes. It is unclear how or why TTX occurs in such a
diverse range of phylogenetically unrelated organisms, however
a popular hypothesis proposes that a symbiotic or commensal
bacterium living within those organisms is responsible for TTX
production.
Analytical TTX detection methods are not a focus of this
review, however, it is important to note that some controversy
(Matsumura, 1995) regarding the specificity of current TTX detection methods exist. No one method of detection, whether it be
chemically (e.g. HPLC) or based on antibody affinity (e.g. ELISA)
has proven unambiguous. In Section 5, we propose that TTX can
be detected using a combination of the above methods as well as
modern molecular techniques, as has been reported for saxitoxin
(Al-Tebrineh et al., 2010).
3.1. Bacterial origins of tetrodotoxin
Bacterial symbiosis, that is, bacteria living within other organisms conferring a positive life sustaining interaction, is a common
occurrence in marine animals. Many symbiotic bacteria are located
in specific organs within their host and some function to benefit
the host via the production of secondary metabolites that provide a means of chemical defense. The conditions within these
organs are tightly regulated to provide optimum conditions for
symbiont growth and metabolite production. For example, host
organisms may produce small signaling molecules that induce
secondary metabolite production (Demain, 1998). The lack of or
absence altogether of inducers derived from the host, may account
for the lower levels of secondary metabolite production achieved
in laboratory cultures. This is important to note in the context of
the biosynthesis studies to be discussed later. We now know, and
many examples exist to support this, that some natural products
originally attributed to macroorganisms have been identified from
cultured microbial symbionts, in particular those associated with
marine animals. Marine sponges (phylum: Porifera) are an important source of bioactive compounds. In recent years, many of these
compounds have been shown to be produced by bacteria living
in a symbiotic relationship to sponge (Hill, 2004). For example, the
bioactive compounds, theopalauamide and swinholide A, originally
isolated from the Philippines sponge, Theonella swinhoei, have been
found to be associated with filamentous and unicellular symbiotic bacteria, respectively (Bewley et al., 1996). Further evidence
is provided by the symbiotic relationship between nematodes and
species of the bacterium genus Xenorhabdus. Xenorhabdus spp.
reside within specialized vesicles in juvenile nematodes and are
capable of producing insecticidal “Tc toxins” (Ffrench-Constant and
Bowen, 2000). When juvenile nematodes are ingested by insects,
the bacterium is excreted from the host and these bacteria then
release Tc toxin which kills the insect. The dead animal is then used
as a rich source of nutrients and a habitat for nematode reproduction (Ffrench-Constant and Bowen, 2000).
The widespread occurrence of TTX in phylogenetically distinct
organisms (Table 2) strongly suggests that symbiotic bacteria play a
role in TTX biosynthesis. Horizontal gene transfer between higher
animals has yet to be documented, and convergent evolution of
TTX biosynthesis genes in such a large number of organisms is
R. Chau et al. / Aquatic Toxicology 104 (2011) 61–72
65
Table 3
Distribution of TTX-producing bacteria.
TTX producing bacteria
Source of bacterial isolate
Reference
Vibrio spp.
Atergatis floridus (reef crab)
Fugu vermicularis vermicularis (common puffer)
Four species of Chaetognaths (arrowworms)
Astropecten polyacanthus (comb seastar)
Hapalochlaena maculosa (blue-ringed octopus)
Deep sea sediment
Fugu vermicularis radialis (common puffer)
Seven species of nemertean worms
Puffer fish species
Nassarius semiplicatus (sea snail)
Niotha clathrata (marine gastropod)
Noguchi et al. (1983)
Noguchi et al. (1987)
Thuesen and Kogure (1989)
Narita et al. (1987)
Hwang et al. (1989)
Do et al. (1990)
Lee et al. (2000)
Carroll et al. (2003)
Yu et al. (2004)
Wang et al. (2008)
Cheng et al. (1995)
Pseudomonas spp.
Jania spp. (red alga)
Fugu poecilonotus (common puffer)
Hapalochlaena maculosa
Niotha clathrata
Yasumoto et al. (1986)
Yotsu et al. (1987)
Hwang et al. (1989)
Cheng et al. (1995)
Bacillus spp.
Hapalochlaena maculosa
Deep sea sediment
Freshwater sediment
Fugu rubripes (common puffer)
Hwang et al. (1989)
Do et al. (1990)
Do et al. (1993)
Wu et al. (2005a,b)
Alteromonas spp.
Hapalochlaena maculosa
Deep sea sediment
Hwang et al. (1989)
Do et al. (1990)
Aeromonas spp.
Deep sea sediment
Niotha clathrata
Do et al. (1990)
Cheng et al. (1995)
Micrococcus spp.
Deep sea sediment
Freshwater sediment
Do et al. (1990)
Do et al. (1993)
Pseudoalteromonas spp.
Meoma ventricosa (sea urchin)
Ritchie et al. (2000)
Serratia marcescens
Puffer fish species
Yan et al. (2005, 2004)
Shewanella putrefaciens
Takifugu niphobles (common puffer)
Matsui et al. (1989)
Acinetobacter spp.
Streptomyces spp.
Caulobacter spp.
Flavobacterium spp.
Plesiomonas spp.
Microbacterium arabinogalactanolyticum
Nocardiopsis dassonvillei
Actinomycete spp.
Marinomonas spp.
Tenacibaculum spp.
Deep sea sediment
Marine sediment
Freshwater sediment
Freshwater sediment
Niotha clathrata
Marine Puffer fish
Fugu rubripes
Fugu rubripes
Nassarius semiplicatus
Nassarius semiplicatus
Do et al. (1990)
Do et al. (1991)
Do et al. (1993)
Do et al. (1993)
Cheng et al. (1995)
Yu et al. (2004)
Wu et al. (2005a,b)
Wu et al. (2005a,b)
Wang et al. (2008)
Wang et al. (2008)
unlikely. Further, direct evidence of a bacterial origin is given by
the many TTX-producing bacterial species isolated from animals
(Table 3). TTX has been isolated from several species of puffer fish
(Table 2). Puffer fish are one of the most extensively studied organisms for the purpose of determining the origin of TTX and indeed
TTX-producing bacteria have also been isolated from these species.
Interestingly, these fish lose the ability to produce TTX when raised
in captivity (Noguchi et al., 2006a,b), supporting the possibility that
TTX is bio-accumulated through the food chain. The amount of TTX
isolated from laboratory cultures is usually quite small when compared to the high levels of TTX usually found in host animals lending
further support to the bio-accumulation hypothesis (Noguchi and
Arakawa, 2008), however, the counter-argument that these bacteria require specific inducers to promote TTX production could
also be valid in some cases (Proksch et al., 2002). Further to this,
the possibility that unculturable microorganisms play a role in TTX
biosynthesis cannot be ignored. It is estimated that less than 1%
of bacteria within a particular community are culturable, thus if
unculturable bacteria synthesize TTX precursors, or produce chemical inducers of TTX biosynthesis, production of the toxin in culture
will be reduced.
TTX-producing bacteria have been isolated from most but not
all animals containing TTX. Hence, there is still dispute regarding
TTX biosynthesis by bacteria. Particularly in newt species, which
have shown no evidence of TTX producing bacteria (Lehman and
Brodie, 2004). A bacterial origin of TTX could ultimately be proven
by understanding the biosynthetic pathways involved in its assembly and also the genes regulating TTX biosynthesis. The proposed
mechanisms of TTX-assembly in bacteria will be discussed later in
this review.
3.2. Non-bacterial origins of tetrodotoxin
A novel neurotoxin, initially classified as tarichatoxin was isolated from the newt, Taricha torosa, in 1963 (Brown and Mosher,
1963). Advances in structural understanding of both tarichatoxin
and TTX in 1964 revealed that both toxins were identical (Mosher
et al., 1964). Henceforth, the name tetrodotoxin was used to classify
this toxin.
Although bacteria have been found to be present in many TTXproducing animals, evidence has suggested that such a symbiotic
relationship does not exist in newts. In Taricha granulosa, tissues
with the highest levels of TTX include skin, ovaries, and muscle
(Wakely et al., 1966). However, an attempt to investigate the bacterial population within T. granulosa, using PCR methods showed that
these tissues did not contain any bacterial DNA except in gastrointestinal tissue, which had low amounts of TTX (Lehman and Brodie,
2004). Hence, a conclusion was made that newts were unlikely to
harbor TTX-producing bacteria. The study did not account for the
immensely greater proportion of newt to bacterial DNA, which may
66
R. Chau et al. / Aquatic Toxicology 104 (2011) 61–72
Fig. 3. Shimizu and co-worker’s proposed biosyntheses of tetrodotoxin (Kotaki and Shimizu, 1993).
have likely interfered with PCR amplification. Although low levels
of TTX are found in the gastrointestinal tract of newts, the presence
of TTX-producing bacteria in these tissues cannot be discounted at
this stage.
3.3. Dietary origins of tetrodotoxin in higher animals
Tetrodotoxin-producing bacteria have been isolated from both
marine and freshwater sediments. It has been proposed that these
bacteria are then ingested by lower-order organisms, which are
in turn eaten by larger predators, biomagnifying TTX in the food
chain (Noguchi and Arakawa, 2008; Noguchi et al., 2006b). It is
well known that wild puffer fish contain TTX, however studies
have shown that captured puffer fish (Takifugu rubripes) raised
for three years in controlled, filtered seawater aquaria and fed a
non-TTX containing diet lose toxicity over time (Noguchi et al.,
2006a). TTX toxicity was assessed using a mouse bioassay, and
the fish were eventually found to be non-toxic (Noguchi et al.,
2006a). The authors thus proposed that TTX in the puffer fish was
derived from an exogenous source rather than a symbiotic or commensal microorganism. In another study, non-toxic, cultured Fugu
niphobles were fed TTX-containing diets for 30 days. Using liquid chromatography-fluorescence detection (LC-FLD) to analyse for
TTX (Yasumoto and Michishita, 1985) it was found that the toxicity of these fishes was restored to a level of 120 ␮g in the liver. The
toxicity of F. niphobles slowly dropped to a level of 50 ␮g in the liver
that was then maintained for 200 days (Kono et al., 2008).
Examples such as these are not uncommon in the marine environment, and a similar mechanism has been proposed for the
observed bioaccumulation of the protein phosphatase inhibitor
okadaic acid in marine sponges. Okadaic acid (OA) which was
originally isolated from the marine sponge Halichondria okadai
(Tachibana et al., 1981) was later shown to be produced in a culture
of the benthic dinoflagellate Prorocentrum lima (Murakami et al.,
1982). Whether bioaccumulation of OA in the sponge results from
P. lima living in the sponge as a symbiont or from settling on the
sponge is debatable.
4. Biosynthesis of tetrodotoxin
4.1. Feeding experiments
Feeding studies with labelled precursors have traditionally been
used to unravel a molecule’s biosynthesis. Prior to the advent of
high field NMR, detection using autoradiography and mass spectrometry of radioactive isotopes was used to measure precursor
incorporation. The use of NMR as an analytical tool not only allowed
for the quantification of labelled substrate incorporation but also
the location of labels on the molecule and thus the ultimate fate
of the precursors. An animal feeding study conducted in the early
1980s (Shimizu and Kobayashi, 1983) is the only significant study
reported with the aim of elucidating the TTX biosynthetic pathway.
To the best of our knowledge, no feeding study with respect to TTX
biosynthesis has been conducted using bacterial cultures.
Shimizu’s feeding protocol was based on two proposed biosyntheses of the TTX carbon backbone as displayed in Fig. 3. In the next
section, specific structural features of TTX will be highlighted, and
hypotheses into its origin discussed.
Toxic newts, Taricha torosa and Taricha granuloa, were fed
a diet containing either [2-14 C]-acetate, [guanido-14 C]-arginine,
[ureido-14 C]-citrulline or [U-14 C]-glucose. Although radioactivity
was detected in primary metabolites, such as cholesterol and some
amides, none of the TTX produced by the newts was labelled.
Subsequent experiments introducing the labelled substrates via
injection, and total immersion of newts in the substrates also
yielded negative results. Even though the feeding experiments
could not confirm incorporation of any of the substrates into TTX
via the proposed biosynthesis pathway, it did not completely eliminate the possibility that TTX was assembled from these proposed
biosynthetic building blocks.
If a bacterial origin was present in newts, then an oral route
may not have properly introduced the labelled substrates to the site
of bacterial TTX biosynthesis. Furthermore, these substrates may
be utilized in the newt’s primary metabolism before they reached
the bacterial site. This may explain the incorporation of labelledsubstrates into primary metabolites derived from cholesterol and
amino acid synthesis pathways (Shimizu and Kobayashi, 1983).
It has also been proposed that TTX precursors may not be synthesized de novo in Taricha spp., but rather acquired through their
diet, similar to puffer fish. This could explain the null result in
Shimizu’s study. However, it is now known that newts increase in
toxicity over time when fed a non-TTX containing diet (Hanifin and
Brodie, 2002), whereas puffer fish decrease in toxicity over time
(Noguchi et al., 2006a). This suggests that the biogenesis of TTX
in newts may be different to other organisms, and may have an
endogenous rather than a dietary origin.
4.2. The genetic basis for a proposed tetrodotoxin biosynthesis
pathway
At present the biosynthesis of TTX is largely unknown, with only
a few proposed pathways to TTX proposed (Fig. 3), and there is no
convincing data in the literature to support any of these hypotheses.
Its unique structure and thus the lack of comparative biosynthesis studies have hindered the development of molecular tools for
the purpose of unraveling the pathway. Our efforts toward understanding the biosynthesis of TTX and the genetics underpinning
its assembly are based upon the notion that the genes coding the
biosynthesis enzymes are clustered on the producing organism’s
genome, therefore the positive identification of one proposed coding gene or region could allow for the identification of the complete
gene cluster. Due mainly to its unique structure, there are several
plausible routes to, and many possible enzymes involved in, TTX
biosynthesis. It is likely that some biosynthesis enzymes involved
R. Chau et al. / Aquatic Toxicology 104 (2011) 61–72
67
Fig. 4. Examples of amidinotransfer in the biosynthesis of cylindrospermopsin (A), saxitoxin (B) and as proposed for tetrodotoxin (C).
in the assembly of similar structural moieties could, by analogy, be
functioning in a similar manner in a TTX producer. Some possible
TTX biosynthesis enzymes will be discussed in terms of the structural fragments incorporated into the final product in the following
sections.
4.2.1. The guanidinium moiety of tetrodotoxin
The guanidinium moiety of TTX (Figs. 2 and 3) is important
for its toxicity and serves as a good target, initially, for predicting the biosynthesis of the molecule due to its rarity in secondary
metabolites. When binding to the voltage-gated sodium channels,
the guanidinium moiety forms a salt bridge between the hydroxyl
groups on the sodium channels (Lee and Ruben, 2008; Lipkind and
Fozzard, 2005) and thus is vital for correct binding of TTX to the
receptor.
Two routes to the incorporation of a guanidinium moiety in
TTX are plausible. The moiety could be either transfered from an
amidino-donor via an amidinotransferase, or incorporated via a
non-ribosomal peptide synthetase module incorporating arginine.
Both of these routes have parallels in characterized natural product
biosyntheses, and are equally likely to occur. Apart from the feeding
study mentioned previously (Shimizu and Kobayashi, 1983) there
has been virtually no research into the origin of the guanidinium
moiety of TTX.
4.2.1.1. Amidinotransferase involvement in tetrodotoxin biosynthesis. Amidinotransferases are enzymes which are responsible for the
transfer of guanidino groups from donors to acceptors. A proposed
origin of the guanidinium moiety in TTX is via amidinotransfer from
an amidino donor such as arginine to a starter substrate in TTX
biosynthesis (Fig. 4). These enzymes are involved in the biosynthesis of other toxins including phaseolotoxin (Hernández-Guzmán
and Alvarez-Morales, 2001; Märkisch and Reuter, 1990), saxitoxin
(Kellmann et al., 2008) and cylindrospermopsin (Mihali et al., 2008)
(Fig. 1).
Phaseolotoxin is a phytotoxin produced solely by the bacteria
Pseudomonas syringae pv. Phaseolicola. Although the biosynthesis
of this toxin has yet to be fully elucidated, amidinotransferase
activity has been found to be directly linked to toxin biosynthesis (Hernández-Guzmán and Alvarez-Morales, 2001; Märkisch
and Reuter, 1990). To determine the substrates and product of
the amidinotransferase, [guanido-14 C]-arginine was successfully
used as a guanidino-donor and was transferred to lysine which
functions as the acceptor, to produce labelled homoarginine.
When repeated with unlabelled arginine coupled with [U-14 C]lysine as a guanidino-acceptor a similar result was reported. All
other donor–acceptor combinations yielded very low levels of
labelled product, or a product which did not fit the biosynthesis
of phaseolotoxin. Hence, phaseolotoxin biosynthesis involves an
amidinotransfer from l-arginine to l-lysine forming homoarginine
and ornithine which are substrates for later steps in the phaseolotoxin biosynthesis pathway (Märkisch and Reuter, 1990).
Cylindrospermopsin is a neurotoxin produced by several species
of cyanobacteria including, Cylindrospermopsis raciborskii, Aphanizomenon ovalisporum and Aphanizomenon flos-aquae (Banker et al.,
1997; Bourke et al., 1983). The cylindrospermopsin biosynthesis gene cluster has recently been sequenced in C. raciborskii
(Mihali et al., 2008). In the proposed pathway, the amidinotransferase CyrA catalyzes the transfer of an amidino group from
l-arginine donor to the acceptor l-glycine, forming guanidinoacetate, which is then incorporated into subsequent biosynthesis
steps (Mihali et al., 2008) (Fig. 4). As in cylindrospermopsin biosynthesis, saxitoxin uses the analogous amidinotransferase SxtG in its
biosynthesis. Saxitoxin shares many similarities with TTX, they are
both relatively small toxins, saxitoxin (C10 H17 N7 O4 ) is 299 g mol−1 ,
and TTX (C11 H17 N3 O8 ) is 319 g mol−1 , both possess a guanidino
group, and both are sodium channel blockers which bind to the
same site on voltage-gated sodium channels (Lipkind and Fozzard,
1994). Importantly, both have been proven to be biosynthesised
by microorganisms. Interestingly, many animals such as xanthid
68
R. Chau et al. / Aquatic Toxicology 104 (2011) 61–72
Fig. 5. Proposed biosynthesis of the caged, type II PKS product, TW93h in Streptomyces coelicolor (A). DMAPP derived isoprene incorporation into zeatin (B), and the
analogous incorporation of an IPP derived isoprene into TTX (C) as proposed by Shimizu (Kotaki and Shimizu, 1993).
crabs, Atergatis floridus, and puffer fish, Takifugu oblongus and Fugu
pardalis, which are thought of as TTX producers have also been
linked to saxitoxin production (Arakawa et al., 1995; Jang and
Yotsu-Yamashita, 2006; Ngy et al., 2009). It is plausible that the
biosynthesis of saxitoxin and tetrodotoxin may involve similar
mechanisms.
A biosynthesis pathway for saxitoxin has been proposed in
C. raciborskii, involving an amidinotransferase which functions to
transfer the amidino group from l-arginine to the saxitoxin precursor 4,7-diguanidino-3-oxoheptane (Kellmann et al., 2008) (Fig. 4).
The amidinotransferase sequence showed high amino acid similarity to the related l-arginine:l-lysine amidinotransferases and are
similar to the well-characterised amidinotransferase enzyme from
P. syringae pv. Phaseolicola, and Streptomyces spp. involved in phaseolotoxin biosynthesis (Hernández-Guzmán and Alvarez-Morales,
2001).
In both of these biosynthesis pathways, the gene encoding the
amidinotransferase is clustered with other biosynthesis enzymes
including non-ribosomal peptide synthetases (NRPS) and polyketide synthases (PKS). We suspect that the guanidinium moiety of
TTX is derived in a similar manner to the toxins discussed above.
In a proposed TTX biosynthesis, an amidinotransferase could facil-
itate the attachment of an amidino group onto ␤-alanine to form a
guanidinopropionate. This substrate could then be utilized as the
precursor for subsequent biosynthesis steps, possibly encoded by
PKS genes.
4.2.1.2. Non-ribosomal peptide synthetase involvement in
tetrodotoxin biosynthesis. The guanidinium moiety of TTX may be
derived from arginine as originally proposed by Shimizu (Kotaki
and Shimizu, 1993). Arginine is proposed to condense with either
a branched apiose sugar or an isoprene unit to form the general
structure of TTX (Figs. 3 and 5). The condensation of arginine in
this manner would likely require the use of peptide biosynthesis
machinery. Ribosomal peptide biosynthesis is responsible for the
production of a large diversity of natural products. These products
consist of post-translationally modified polypeptides that have
been recently reviewed (McIntosh et al., 2009). However, such
polypeptide pathways are difficult to envision in the structure of
TTX and thus a NRPS-type assembly is more likely.
NRPSs and PKSs are multifunctional enzyme complexes that
sequentially assemble amino acid and small carboxylate derived
precursor building blocks, respectively, into their products in an
assembly line-like fashion. Both NRPSs and type I PKSs share
R. Chau et al. / Aquatic Toxicology 104 (2011) 61–72
similar architectures with their respective modules containing a
minimum of three enzyme domains. NRPS modules contain an ATPdependent adenylation (A) domain which activates a specific or
preferred amino acid, a peptidyl carrier protein (PCP) for tethering substrates during the assembly, and a condensation domain (C)
which catalyzes the formation of amide bonds between PCP-bound
amino acids.
An alternate route to the incorporation of an arginine-derived
guanidinium could be via an NRPS. These enzymatic systems use
amino acid substrates in secondary metabolite biosynthesis. Unlike
ribosomal peptide synthases, NRPSs are able to incorporate nonproteinogenic, that is modified or unusual, amino acids and are
commonly associated with other biosynthesis enzymatic systems
including PKSs (Cane and Walsh, 1999). Combinations of such genes
allows for the highly programmed biosynthesis of many unique
compounds, and incorporate one or more amino acids. PKS modules
contain an acyl transferase (AT) which selects a preferred acyl-CoA
thioester substrate, acyl carrier protein (ACP) and a ketosynthase
(KS) which catalyzes the condensation of two ACP-bound substrates. In both systems, each module extends the backbone of the
molecule by one unit. These modules often contain a number of
additional catalytic domains which function to tailor the assembling molecule and therefore generate further structural diversity.
The assembled molecule is then released from the enzyme complex,
usually by a thioesterase (TE) domain which may also function to
direct cyclization of the final product. A recent review by Walsh
and co-workers should be consulted for an in depth discussion of
the assembly of NRPS and PKS derived products (Sattely and Walsh,
2008).
Such hybrid PKS-NRPS systems are involved in the biosynthesis
of cytochalasin G, produced by fungal isolates including Zygosporium masonii (Schumann and Hertweck, 2007) and aspyridone,
produced by Aspergillus nidulans (Bergmann et al., 2007) (Fig. 1).
Both of these molecules are examples of natural products incorporating a single amino acid and a PKS derived moiety. In the
biosyntheses of cytochalasan and aspyridone, tryptophan and tyrosine, respectively, are incorporated into a growing enzyme bound
polyketide chain prior to thioesterase (TE) mediated release and
cyclization. A similar mechanism of arginine condensation catalyzed by an NRPS is plausible in TTX biosynthesis and in such a
pathway, the NRPS would likely be clustered with other biosynthesis enzymes to assemble the carbon backbone, which may be
polyketide, sugar or terpene derived as previously proposed (Kotaki
and Shimizu, 1993; Woodward and Gougoutas, 1964; Yasumoto
et al., 1988). The proposed biosynthesis of the molecule, however,
would require further experimental evidence to support a seemingly unusual and unprecedented assembly.
69
there are no biosynthetic precedents in the literature. Moore and
co-workers have reported a polyketide, TW93h, containing a 2,4dioxaadamantane structure (Fig. 5), similar to that of TTX indicating
that such structures are possible via a PKS pathway (Shen et al.,
1999). It is notable that TW93h, however, is assembled by a type
II PKS rather than the above mentioned type I PKS and thus such
systems should not be ignored in examining the TTX biosynthesis
pathway. Examples of hybrid NRPS/type II PKSs, however, have not
been reported.
4.2.2.2. A sugar derived carbon backbone fragment. Shimizu proposed that the highly oxygenated structure of TTX may be derived
via the condensation of arginine with an oxygenated, branched C5
sugar such as apiose, which is known to occur in marine environments (Kotaki and Shimizu, 1993). Apiose incorporation into
natural products is rare and only plant natural products such as
conyzasaponin, produced by Conyza blinii (Su et al., 2001) (Fig. 1)
have been reported to possess apiose sugars. However, the identification of deoxy-TTX precursors indicates that apiose incorporation
is unlikely. The incorporation of apiose into TTX would indeed
result in a highly oxygenated carbon backbone, such as that found in
TTX, however, the isolation of deoxy-TTX indicates that a less oxygenated substrate, such as an isoprene unit rather than an apiose
sugar is more likely (Kotaki and Shimizu, 1993).
4.2.2.3. A terpene derived carbon backbone fragment. Natural products of the terpene class are derived from C5 building blocks
such as dimethylallyl pyrophosphate (DMAPP) and isopentenyl
diphosphate (IPP) (Dewick, 2002). Meroterpenes are a class of
natural product incorporating isoprene units and other moieties derived from different pathways, including polyketides.
Zeatin, a DMAPP derived plant hormone is biosynthesized via
an N-prenylation of adenosine mono-phosphate, catalyzed by the
enzyme adenosine phosphate-isopentenyltransferase (KamadaNobusada and Sakakibara, 2009), to form the zeatin precursor
N6 -(2 -isopentenyl)adenine riboside phosphate (Fig. 5). Likewise,
Yasumoto proposed that a mixed pathway incorporating an isoprene unit and a single arginine to form TTX could be plausible
(Yasumoto et al., 1988). Stepwise oxygenation reactions would
then follow to finally yield the highly oxygenated carbon backbone
present in TTX (Fig. 5). There are many characterized enzymes that
catalyze isoprene attachment, however, an enzyme for the prenylation of amino acids has yet to be reported. Regardless of whether
TTX biosynthesis incorporates a polyketide, sugar or terpene substrate, the pathway employed would be unique, and unlike any
other pathway investigated to date.
5. Future directions
4.2.2. The carbon backbone of tetrodotoxin
TTX
contains
a
unique,
highly
oxygenated,
2,4dioxaadamantane carbon backbone. Due to its unique structure,
many hypotheses regarding the origin of the carbon backbone of
TTX have been proposed, including polyketide (Woodward and
Gougoutas, 1964), C5 branched sugar (Kotaki and Shimizu, 1993),
and C5 isoprene origins (Yasumoto et al., 1988). Incorporation
of C5 units would require priming by arginine rather than an
amidinotransferase facilitated guanidine transfer to account for all
carbon atoms in TTX.
4.2.2.1. A polyketide derived carbon backbone. A polyketide pathway as proposed by Woodward and Gougoutas (1964) is feasible
if indeed the guanidinium is derived from an amidinotransfer onto
a three carbon unit acceptor (Fig. 5) which is then extended by
malonyl-CoA derived acetates, to account for all 11 carbons in
the backbone (Fig. 5). Conversion of the nascent polyketide into
the caged structure of TTX does not appear straight forward and
Despite its long history and a thorough knowledge of its toxicity
and pharmacology, neither the pathway to TTX nor even the biogenic origin of TTX is known. The debate into whether TTX is derived
from bacteria or is endogenous to the host animals is on-going and
the only published study into the substrates of TTX biosynthesis
proved inconclusive.
The advancement of molecular techniques has led to a better understanding of small molecule biosynthesis pathways at a
genetic level. Today, the elucidation of biosynthesis gene clusters
via genetics-based methods has allowed for the partial or complete
assembly of molecules with unusual structures to be proposed.
In general, our working knowledge of biosynthesis gene clusters
allows not only for the identification of the catalytic enzymes but
also a particular enzyme’s preferred substrate and co-factor. Structural similarities between small natural product molecules, which
include toxins, more often than not suggest a common biosynthetic
route or precursor. Some natural products which are structurally
70
R. Chau et al. / Aquatic Toxicology 104 (2011) 61–72
related in part to TTX have been highlighted in this review and
their biosynthesis involves enzymes which may also utilized in TTX
assembly.
We propose that the use of genetic methods, specifically, targeting genes coding for biosynthesis enzymes is the way forward
to solving the mystery of TTX assembly and its origin. This is based
on the notion that the biosyntheses of many toxins utilize enzymes
whose mechanisms are likely paralleled in TTX assembly. Screening
microorganisms implicated in TTX production for such biosynthesis genes using PCR-based methods may reveal the unique enzymes
responsible for TTX production and, coupled with traditional feeding experiments, may finally lead to the elucidation of the TTX
pathway.
Unculturable or yet-to-be cultured microbes have been largely
ignored as a focus of biosynthesis studies. The development of
metagenomic sequencing technologies, and heterologous expression systems, has allowed the biosynthesis gene clusters from
unculturable bacteria to be further investigated. Interrogating environmental DNA libraries has provided access to new biosynthesis
pathways from unculturable microorganisms. Such metagenomic
approaches should also be considered in order to help understand
TTX’s assembly.
Although this review has attempted to highlight possible
biosynthesis routes to TTX based on current knowledge, investigation into its biosynthesis has proven difficult due to the novelty
of its structure with many studies rendering null results. A combined approach using both traditional and modern methods will
in our opinion shed additional light on this problem. We are certain that a complete understanding of TTXs assembly will reveal
many novel biosynthetic mechanisms and thus novel biosynthesis
enzymes which maybe of commercial use. Preliminary investigations in our laboratory have revealed a number of candidate
symbiotic microorganisms from TTX sources which appear to possess analogues of genes discussed in this review, which may be
involved in TTX biosynthesis.
References
Akopian, A.N., Souslova, V., England, S., Okuse, K., Ogata, N., Ure, J., Smith, A., Kerr, B.J.,
McMahon, S.B., Boyce, S., Hill, R., Stanfa, L.C., Dickenson, A.H., Wood, J.N., 1999.
The tetrodotoxin-resistant sodium channel SNS has a specialized function in
pain pathways. Nat. Neurosci. 2, 541–548.
Al-Tebrineh, J., Mihali, T.K., Pomati, F., Neilan, B.A., 2010. Detection of saxitoxinproducing cyanobacteria and Anabaena circinalis in environmental water blooms
by quantitative PCR. Appl. Environ. Microbiol. 76, 7836–7842.
Arakawa, O., Nishio, S., Noguchi, T., Shida, Y., Onoue, Y., 1995. A new saxitoxin
analogue from a xanthid crab Atergatis floridus. Toxicon 33, 1577–1584.
Bacher, A., Rieder, C., Eichinger, D., Arigoni, D., Fuchs, G., Eisenreich, W., 1998.
Elucidation of novel biosynthetic pathways and metabolite flux patterns by
retrobiosynthetic NMR analysis. FEMS Microbiol. Rev. 22, 567–598.
Banker, R., Carmeli, S., Hadas, O., Teltsch, B., Porat, R., Sukenik, A., 1997. Identification
of cylindrospermopsin in Aphanizomenon ovalisporum (Cyanophyceae) isolated
from Lake Kinneret, Israel. J. Phycol. 33, 613–616.
Bergmann, S., Schumann, J., Scherlach, K., Lange, C., Brakhage, A.A., Hertweck,
C., 2007. Genomics-driven discovery of PKS-NRPS hybrid metabolites from
Aspergillus nidulans. Nat. Chem. Biol. 3, 213–217.
Bewley, C., Holland, N., Faulkner, D., 1996. Two classes of metabolites from Theonella
swinhoei are localized in distinct populations of bacterial symbionts. Cell. Mol.
Life Sci. 52, 716–722.
Blunt, J.W., Copp, B.R., Munro, M.H.G., Northcote, P.T., Prinsep, M.R., 2010. Marine
natural products. Nat. Prod. Rep., 165–237.
Bourke, A.T.C., Hawes, R.B., Neilson, A., Stallman, N.D., 1983. An outbreak of hepatoenteritis (the Palm Island mystery disease) possibly caused by algal intoxication.
Toxicon 21, 45–48.
Brown, M.S., Mosher, H.S., 1963. Tarichatoxin: Isolation and purification. Science
140, 295–296.
Cane, D.E., Walsh, C.T., 1999. The parallel and convergent universes of polyketide
synthases and nonribosomal peptide synthetases. Chem. Biol. 6, R319–R325.
Carroll, S., McEvoy, E., Gibson, R., 2003. The production of tetrodotoxin-like substances by nemertean worms in conjunction with bacteria. J. Exp. Mar. Biol.
Ecol. 288, 51–63.
Cheng, C.A., Hwang, D.F., Tsai, Y.H., Chen, H.C., Jeng, S.S., Noguchi, T., Ohwada, K., Hasimoto, K., 1995. Microflora and tetrodotoxin-producing bacteria in a gastropod,
Niotha clathrata. Food Chem. Toxicol. 33, 929–934.
Cook, J., 1775. The Journals of Captain James Cook on his voyages of discovery.
Cambridge University Press, London.
Daly, J., Gusovsky, F., Myers, C., Yotsu-Yamashita, M., Yasumoto, T., 1994. First occurrence of tetrodotoxin in a dendrobatid frog (Colostethus inguinalis), with further
reports for the bufonid genus Atelopus. Toxicon 32, 279.
Dao, H., Takata, Y., Sato, S., Fukuyo, Y., Kodama, M., 2009. Frequent occurrence of the
tetrodotoxin-bearing horseshoe crab Carcinoscorpius rotundicauda in Vietnam.
Fish. Sci. 75, 435–438.
Demain, A., 1998. Induction of microbial secondary metabolism. Int. Microbiol. 1,
259–264.
Dewick, P., 2002. The Mevalonate and Deoxyxylulose Phosphate Pathways: Terpenoids and Steroids, Medicinal Natural Products: A Biosynthetic Approach.
John Wiley & Sons Inc, pp. 167–285.
Do, H., Hamasaki, K., Ohwada, K., Simidu, U., Noguchi, T., Shida, Y., Kogure, K., 1993.
Presence of tetrodotoxin and tetrodotoxin-producing bacteria in freshwater
sediments. Appl. Environ. Microbiol. 59, 3934.
Do, H., Kogure, K., Imada, C., Noguchi, T., Ohwada, K., Simidu, U., 1991. Tetrodotoxin
production of actinomycetes isolated from marine sediment. J. Appl. Microbiol.
70, 464–468.
Do, H., Kogure, K., Simidu, U., 1990. Identification of deep-sea-sediment bacteria
which produce tetrodotoxin. Appl. Environ. Microbiol. 56, 1162.
Epstein-Barash, H., Shichor, I., Kwon, A.H., Hall, S., Lawlor, M.W., Langer, R., Kohane,
D.S., 2009. Prolonged duration local anesthesia with minimal toxicity. Proc. Natl.
Acad. Sci. U. S. A. 106, 7125–7130.
Ffrench-Constant, R., Bowen, D., 2000. Novel insecticidal toxins from nematodesymbiotic bacteria. Cell. Mol. Life Sci. 57, 828–833.
Fuhrman, F., 1986. Tetrodotoxin, tarichatoxin, and chiriquitoxin: Historical perspectives. Ann. N. Y. Acad. Sci. 479, 1–14.
Glaser, K.B., Mayer, A.M.S., 2009. A renaissance in marine pharmacology: From preclinical curiosity to clinical reality. Biochem. Pharmacol. 78, 440–448.
Goto, T., Kishi, Y., Takahashi, S., Hirata, Y., 1965. Tetrodotoxin. Tetrahedron 21,
2059–2088.
Hagen, N.A., du Souich, P., Lapointe, B., Ong-Lam, M., Dubuc, B., Walde, D., Love, R.,
Ngoc, A.H., 2008. Tetrodotoxin for moderate to severe cancer pain: a randomized, double blind, parallel design multicenter study. J. Pain Symptom Manage.
35, 420–429.
Hagen, N.A., Fisher, K.M., Lapointe, B., du Souich, P., Chary, S., Moulin, D., Sellers,
E., Ngoc, A.H., 2007. An open-label, multi-dose efficacy and safety study of
intramuscular tetrodotoxin in patients with severe cancer-related pain. J. Pain
Symptom Manage. 34, 171–182.
Halstead, B., 1958. Poisonous fishes. Public Health Rep. 73, 302.
Hanifin, C.T., Brodie, E.D., 2002. Tetrodotoxin levels of the rough-skin newt, Taricha
granulosa, increase in long-term captivity. Toxicon 40, 1149–1153.
Hay, M., Fenical, W., 1996. Chemical ecology and marine biodiversity: Insights and
products from the sea. Oceanography 9, 10–20.
Hernández-Guzmán, G., Alvarez-Morales, A., 2001. Isolation and characterization
of the gene coding for the amidinotransferase involved in the biosynthesis of
phaseolotoxin in Pseudomonas syringae pv. phaseolicola. Mol. Plant Microbe In.
14, 545–554.
Hill, R.T., 2004. Microbes from marine sponges: A treasure trove of biodiversity
for natural products discovery. In: Microbial Diversity and Bioprospecting. ASM
Press, Washington, DC, pp. 177–190.
Hinman, A., Du Bois, J., 2003. A stereoselective synthesis of (−)-tetrodotoxin. J. Am.
Chem. Soc. 125, 11510–11511.
Hwang, D., Arakawa, O., Saito, T., Noguchi, T., Simidu, U., Tsukamoto, K., Shida,
Y., Hashimoto, K., 1989. Tetrodotoxin-producing bacteria from the blue-ringed
octopus Octopus maculosus. Mar. Biol. 100, 327–332.
Hwang, D., Lu, S., Jeng, S., 1991. Occurrence of tetrodotoxin in the gastropods Rapana
rapiformis and R. venosa venosa. Mar. Biol. 111, 65–69.
Jang, J., Yotsu-Yamashita, M., 2006. Distribution of tetrodotoxin, saxitoxin, and
their analogs among tissues of the puffer fish Fugu pardalis. Toxicon 48, 980–
987.
Jeon, J., Narita, H., Nara, M., Noguchi, T., Maruyama, J., Hashimoto, K., 1984. Occurrence of tetrodotoxin in a gastropod mollusk, “araregai” Niotha clathrata. Bull.
Jpn. Soc. Sci. Fish. 50, 2099–2102.
Kalaitzis, J.A., Chau, R., Kohli, G.S., Murray, S.A., Neilan, B.A., 2010. Biosynthesis of
toxic naturally-occurring seafood contaminants. Toxicon 56, 244–258.
Kamada-Nobusada, T., Sakakibara, H., 2009. Molecular basis for cytokinin biosynthesis. Phytochemistry 70, 444–449.
Kellmann, R., Mihali, T., Jeon, Y., Pickford, R., Pomati, F., Neilan, B., 2008. Biosynthetic
intermediate analysis and functional homology reveal a saxitoxin gene cluster
in cyanobacteria. Appl. Environ. Microbiol. 74, 4044.
Kishi, Y., Aratani, M., Fukuyama, T., Nakatsubo, F., Goto, T., Inoue, S., Tanino, H.,
Sugiura, S., Kakoi, H., 1972a. Synthetic studies on tetrodotoxin and related compounds. III. Stereospecific synthesis of an equivalent of acetylated tetrodamine.
J. Am. Chem. Soc. 94, 9217–9219.
Kishi, Y., Fukuyama, T., Aratani, M., Nakatsubo, F., Goto, T., Inoue, S., Tanino, H.,
Sugiura, S., Kakoi, H., 1972b. Synthetic studies on tetrodotoxin and related compounds. IV. Stereospecific total syntheses of DL-tetrodotoxin. J. Am. Chem. Soc.
94, 9219–9221.
Kodama, M., Sato, S., Sakamoto, S., Ogata, T., 1996. Occurrence of tetrodotoxin in
Alexandrium tamarense, a causative dinoflagellate of paralytic shellfish poisoning. Toxicon 34, 1101–1105.
Kohane, D.S., Smith, S.E., Louis, D.N., Colombo, G., Ghoroghchian, P., Hunfeld,
N.G.M., Berde, C.B., Langer, R., 2003. Prolonged duration local anesthesia from
tetrodotoxin-enhanced local anesthetic microspheres. Pain 104, 415–421.
R. Chau et al. / Aquatic Toxicology 104 (2011) 61–72
Kono, M., Matsui, T., Furukawa, K., Yotsu-Yamashita, M., Yamamori, K., 2008. Accumulation of tetrodotoxin and 4,9-anhydrotetrodotoxin in cultured juvenile
kusafugu Fugu niphobles by dietary administration of natural toxic komonfugu
Fugu poecilonotus liver. Toxicon 51, 1269–1273.
Kotaki, Y., Shimizu, Y., 1993. 1-Hydroxy-5,11-dideoxytetrodotoxin, the first Nhydroxy and ring-deoxy derivative of tetrodotoxin found in the newt Taricha
granulosa. J. Am. Chem. Soc. 115, 827–830.
Kungsuwan, A., Nagashima, Y., Noguchi, T., Shida, Y., Suvapeepan, S., Suwansakornkul, P., Hashimoto, K., 1987. Tetrodotoxin in the horseshoe crab Carcinoscorpius rotundicauda inhabiting Thailand. Nippon Suisan Gakk. 53, 261–266.
Lee, C., Ruben, P., 2008. Interaction between voltage-gated sodium channels and the
neurotoxin, tetrodotoxin. Channels, 2.
Lee, M., Jeong, D., Kim, W., Kim, H., Kim, C., Park, W., Park, Y., Kim, K., Kim, H., Kim,
D., 2000. A tetrodotoxin-producing Vibrio strain, LM-1, from the puffer fish Fugu
vermicularis radiatus. Appl. Environ. Microbiol. 66, 1698.
Lehman, E.M., Brodie, E.D., 2004. No evidence for an endosymbiotic bacterial origin
of tetrodotoxin in the newt Taricha granulosa. Toxicon 44, 243–249.
Lipkind, G., Fozzard, H., 1994. A structural model of the tetrodotoxin and saxitoxin
binding site of the Na+ channel. Biophys. J. 66, 1–13.
Lipkind, G.M., Fozzard, H.A., 2005. Molecular modeling of local anesthetic drug binding by voltage-gated sodium channels. Mol. Pharmacol. 68, 1611–1622.
Märkisch, U., Reuter, G., 1990. Biosynthesis of homoarginine and ornithine as precursors of the phytoeffector phaseolotoxin by the amidinotransfer from arginine
to lysine catalyzed by an amidinotransferase in Pseudomonas syringae pv. phaseolicola. J. Basic Microb. 30, 425–433.
Maruyama, J., Noguchi, T., Jeon, J., Harada, T., Hashimoto, K., 1984. Occurrence
of tetrodotoxin in the starfish Astropecten latespinosus. Cell. Mol. Life Sci. 40,
1395–1396.
Matsui, T., Taketsugu, S., Kodama, K., Ishii, A., Yamamori, K., Shimizu, C., 1989.
Production of tetrodotoxin by the intestinal bacteria of a puffer fish Takifugu
niphobles. Nippon Suisan Gakk. 55, 2199–2203.
Matsumura, K., 1995. Reexamination of tetrodotoxin production by bacteria. Appl.
Environ. Microbiol. 61, 3468–3470.
McIntosh, J.A., Donia, M.S., Schmidt, E.W., 2009. Ribosomal peptide natural products:
Bridging the ribosomal and nonribosomal worlds. Nat. Prod. Rep. 26, 537–559.
Mebs, D., Schmidt, K., 1989. Occurrence of tetrodotoxin in the frog Atelopus
oxyrhynchus. Toxicon 27, 819.
Mihali, T., Kellmann, R., Muenchhoff, J., Barrow, K., Neilan, B., 2008. Characterization of the gene cluster responsible for cylindrospermopsin biosynthesis. Appl.
Environ. Microbiol. 74, 716.
Mills, A.R., Passmore, R., 1988. Pelagic paralysis. Lancet 331, 161–164.
Minor, F.W., Greathouse, G.A., Shirk, H.G., Schwartz, A.M., Harris, M., 1954. Biosynthesis of C14-specifically labeled cellulose by Acetobacter xylinum. II. From
D-mannitol-1-C14 with and without ethanol. J. Am. Chem. Soc. 76, 5052–5054.
Miyazawa, K., Jeon, J., Maruyama, J., Noguchi, T., Ito, K., Hashimoto, K., 1986. Occurrence of tetrodotoxin in the flatworm Planocera multitentaculata. Toxicon 24,
645–650.
Miyazawa, K., Noguchi, T., Maruyama, J., Jeon, J., Otsuka, M., Hashimoto, K., 1985.
Occurrence of tetrodotoxin in the starfishes Astropecten polyacanthus and A.
scoparius in the Seto Inland Sea. Mar. Biol. 90, 61–64.
Mosher, H.S., Fuhrman, F.A., Buchwald, H.D., Fischer, H.G., 1964. Tarichatoxintetrodotoxin: a potent neurotoxin. Science 144, 1100–1110.
Murakami, Y., Oshima, Y., Yasumoto, T., 1982. Identification of okadaic acid as a toxic
component of a marine dinoflagellate Prorocentrum lima. Bull. Jpn. Soc. Sci. Fish.
48, 69–72.
Nagashima, Y., Tanaka, N., Shimakura, K., Shiomi, K., Shida, Y., 2001. Occurrence of
tetrodotoxin-related substances in the nontoxic puffer Takifugu xanthopterus.
Toxicon 39, 415–418.
Narahashi, T., 2001. Pharmacology of tetrodotoxin. Toxin Rev. 20, 67–84.
Narahashi, T., 2008. Tetrodotoxin—a brief history. Proc. Jpn. Acad., Ser. B, Phys. Biol.
Sci. 84, 147–154.
Narita, H., Matsubara, S., Miwa, N., Akahane, S., Murakami, M., Goto, T., Nara, M.,
Noguchi, T., Saito, T., Shida, Y., 1987. Vibrio alginolyticus, a TTX-producing bacterium isolated from the starfish Astropecten polyacanthus. Nippon Suisan Gakk.
53, 617–621.
Narita, H., Noguchi, T., Maruyama, J., Ueda, Y., Hashimoto, K., Watanabe, Y., Hida,
K., 1981. Occurrence of Tetrodotoxin in a Trumpet Shell, “Boshubora” Charonia
sauliae. Bull. Jpn. Soc. Sci. Fish. 47, 935–941.
Ngy, L., Yu, C., Taniyama, S., Takatani, T., Arakawa, O., 2009. Co-occurrence of
tetrodotoxin and saxitoxin in Cambodian marine pufferfish Takifugu oblongus.
Afr. J. Mar. Sci., 31.
Noguchi, T., Arakawa, O., 2008. Tetrodotoxin—Distribution and accumulation in
aquatic organisms, and cases of Human intoxication. Mar. Drugs 6, 220–242.
Noguchi, T., Arakawa, O., Takatani, T., 2006a. Toxicity of pufferfish Takifugu rubripes
cultured in netcages at sea or aquaria on land. Comp. Biochem. Physiol. Part D
Genom. Proteom. 1, 153–157.
Noguchi, T., Arakawa, O., Takatani, T., 2006b. TTX accumulation in pufferfish. Comp.
Biochem. Physiol. Part D Genom. Proteom. 1, 145–152.
Noguchi, T., Hashimoto, Y., 1973. Isolation of tetrodotoxin from a goby Gobius
criniger. Toxicon 11, 305–307.
Noguchi, T., Hwang, D., Arakawa, O., Sugita, H., Deguchi, Y., Shida, Y., Hashimoto, K.,
1987. Vibrio alginolyticus, a tetrodotoxin-producing bacterium, in the intestines
of the fish Fugu vermicularis vermicularis. Mar. Biol. 94, 625–630.
Noguchi, T., Maruyama, J., Narita, H., Hashimoto, K., 1984. Occurrence of tetrodotoxin
in the gastropod mollusk Tutufa lissostoma (frog shell). Toxicon 22, 219.
71
Noguchi, T., Maruyama, J., Ueda, Y., Hashimoto, K., Harada, T., 1981. Occurrence of
tetrodotoxin in the Japanese ivory shell Babylonia japonica. Bull. Jpn. Soc. Sci.
Fish. 47, 901–913.
Noguchi, T., Uzu, A., Koyama, K., Hashimoto, K., 1983. Occurrence of tetrodotoxin
as the major toxin in xanthid crab Atergatis floridus. Bull. Jpn. Soc. Sci. Fish. 49,
1887–1892.
Ohyabu, N., Nishikawa, T., Isobe, M., 2003. First asymmetric total synthesis of
tetrodotoxin. J. Am. Chem. Soc. 125, 8798–8805.
Pires, O., Sebben, A., Schwartz, E., Largura, S., Bloch, C., Morales, R., Schwartz, C., 2002.
Occurrence of tetrodotoxin and its analogues in the Brazilian frog Brachycephalus
ephippium (Anura: Brachycephalidae). Toxicon 40, 761–766.
Pires, O., Sebben, A., Schwartz, E., Morales, R., Bloch, C., Schwartz, C., 2005. Further
report of the occurrence of tetrodotoxin and new analogues in the Anuran family
Brachycephalidae. Toxicon 45, 73–79.
Proksch, P., Edrada, R., Ebel, R., 2002. Drugs from the seas-current status and microbiological implications. Appl. Microbiol. Biotechnol. 59, 125–134.
Ritchie, K., Nagelkerken, I., James, S., Smith, G., 2000. Environmental microbiology:
A tetrodotoxin-producing marine pathogen. Nature 404, 354.
Sanford, L.D., Yang, L., Liu, X., Tang, X., 2006. Effects of tetrodotoxin (TTX) inactivation
of the central nucleus of the amygdala (CNA) on dark period sleep and activity.
Brain Res. 1084, 80–88.
Sato, K.-i., Akai, S., Shoji, H., Sugita, N., Yoshida, S., Nagai, Y., Suzuki, K., Nakamura,
Y., Kajihara, Y., Funabashi, M., Yoshimura, J., 2008. Stereoselective and efficient
total synthesis of optically active tetrodotoxin from D-glucose. J. Org. Chem. 73,
1234–1242.
Sattely, E., Walsh, C., 2008. Total biosynthesis: In vitro reconstitution of polyketide
and nonribosomal peptide pathways. Nat. Prod. Rep. 25, 757–793.
Schumann, J., Hertweck, C., 2007. Molecular basis of cytochalasan biosynthesis in
fungi: Gene cluster analysis and evidence for the involvement of a PKS-NRPS
hybrid synthase by RNA silencing. J. Am. Chem. Soc. 129, 9564–9565.
Shen, Y., Yoon, P., Yu, T.-W., Floss, H.G., Hopwood, D., Moore, B.S., 1999. Ectopic
expression of the minimal whiE polyketide synthase generates a library of aromatic polyketides of diverse sizes and shapes. Proc. Natl. Acad. Sci. U. S. A. 96,
3622–3627.
Sheumack, D., Howden, M., Spence, I., 1984. Occurrence of a tetrodotoxin-like
compound in the eggs of the venomous blue-ringed octopus (Hapalochlaena
maculosa). Toxicon 22, 811.
Shimizu, Y., Kobayashi, M., 1983. Apparent lack of tetrodotoxin biosynthesis in captured Taricha torosa and Taricha granulosa. Chem. Pharm. Bull. 31, 3625–3631.
Shiu, Y., Lu, Y., Tsai, Y., Chen, S., Hwang, D., 2003. Occurrence of tetrodotoxin in
the causative gastropod Polinices didyma and another gastropod Natica lineata
collected from western Taiwan. J. Food Drug Anal. 11, 159–163.
Su, Y., Koike, K., Guo, D., Satou, T., Liu, J., Zheng, J., Nikaido, T., 2001. New
apiose-containing triterpenoid saponins from Conyza blinii. Tetrahedron 57,
6721–6726.
Tachibana, K., Scheuer, P.J., Tsukitani, Y., Kikuchi, H., Van Engen, D., Clardy, J., Gopichand, Y., Schmitz, F.J., 1981. Okadaic acid, a cytotoxic polyether from two marine
sponges of the genus Halichondria. J. Am. Chem. Soc. 103, 2469–2471.
Thuesen, E., Kogure, K., 1989. Bacterial production of tetrodotoxin in four species of
Chaetognatha. Biol. Bull. 176, 191.
Tsai, Y., Hwang, D., Chai, T., Jeng, S., 1995. Occurrence of tetrodotoxin and paralytic shellfish poison in the Taiwanese crab Lophozozymus pictor. Toxicon 33,
1669–1673.
Tsuda, K., Ikuma, S., Kawamura, M., Tachikawa, R., Sakai, K., 1964. Tetrodotoxin.
VII. On the structure of tetrodotoxin and its derivatives. Chem. Pharm. Bull. 12,
1356–1374.
Udwary, D.W., Zeigler, L., Asolkar, R.N., Singan, V., Lapidus, A., Fenical, W., Jensen, P.R.,
Moore, B.S., 2007. Genome sequencing reveals complex secondary metabolome
in the marine actinomycete Salinispora tropica. Proc. Natl. Acad. Sci. U. S. A. 104,
10376–10381.
Wakely, J.F., Fuhrman, G.J., Fuhrman, F.A., Fischer, H.G., Mosher, H.S., 1966. The occurrence of tetrodotoxin (tarichatoxin) in amphibia and the distribution of the toxin
in the organs of newts (Taricha). Toxicon 3, 195–203.
Wang, X., Yu, R., Luo, X., Zhou, M., Lin, X., 2008. Toxin-screening and identification
of bacteria isolated from highly toxic marine gastropod Nassarius semiplicatus.
Toxicon 52, 55–61.
Woodward, R., Gougoutas, J., 1964. The structure of tetrodotoxin. J. Am. Chem. Soc.
86, 5030–15030.
Wu, Z., Xie, L., Xia, G., Zhang, J., Nie, Y., Hu, J., Wang, S., Zhang, R., 2005a. A new
tetrodotoxin-producing actinomycete, Nocardiopsis dassonvillei, isolated from
the ovaries of puffer fish Fugu rubripes. Toxicon 45, 851–859.
Wu, Z., Yang, Y., Xie, L., Xia, G., Hu, J., Wang, S., Zhang, R., 2005b. Toxicity and distribution of tetrodotoxin-producing bacteria in puffer fish Fugu rubripes collected
from the Bohai Sea of China. Toxicon 46, 471–476.
Yan, Q., Yu, P., Li, H., 2005. Detection of tetrodotoxin and bacterial production by
Serratia marcescens. World J. Microbiol. Biotechnol. 21, 1255–1258.
Yasumoto, T., Michishita, T., 1985. Fluorometric determination of tetrodotoxin by
high performance liquid chromatography. Agric. Biol. Chem. 49, 3077–3080.
Yasumoto, T., Yasumura, D., Yotsu, M., Michishita, T., Endo, A., Kotaki, Y., 1986. Bacterial production of tetrodotoxin and anhydrotetrodotoxin. Agric. Biol. Chem.
50, 793–795.
Yasumoto, T., Yotsu, M., Murata, M., Naoki, H., 1988. New tetrodotoxin analogs from
the newt Cynops ensicauda. J. Am. Chem. Soc. 110, 2344–2345.
Yokoo, A., 1950. Study on chemical purification of tetrodotoxin (3)—purification of
spheroidine. J. Chem. Soc. Japan 71, 590–592.
72
R. Chau et al. / Aquatic Toxicology 104 (2011) 61–72
Yotsu-Yamashita, M., Mebs, D., 2003. Occurrence of 11-oxotetrodotoxin in the redspotted newt, Notophthalmus viridescens, and further studies on the levels of
tetrodotoxin and its analogues in the newt’s efts. Toxicon 41, 893–897.
Yotsu, M., Yamazaki, T., Meguro, Y., Endo, A., Murata, M., Naoki, H., Yasumoto, T.,
1987. Production of tetrodotoxin and its derivatives by Pseudomonas sp. isolated
from the skin of a pufferfish. Toxicon 25, 225.
Yu, C., Yu, P., Chan, P., Yan, Q., Wong, P., 2004. Two novel species of tetrodotoxinproducing bacteria isolated from toxic marine puffer fishes. Toxicon 44,
641–647.
Zhou, M., Shum, F., 2003. United States Patent: Method of extracting tetrodotoxin.
In: Office, U.S.P. (Ed.). Wex Medical Instrumtation Co., Ltd., Hong Kong, p. 22.