The chemistry and chemical ecology of nudibranchs

Contents

• 1 Introduction

• 2 Taxonomy

• 3 The origin of nudibranch natural products

• 4 Scope of review

• 5 Dorid nudibranchs
  • 5.1 Bathydoridoidea
  • 5.2 Doridoidea
  • 5.3 Onchidoridoidea
  • 5.4 Phyllidioidea
  • 5.5 Polyceroidea

• 6 Cladobranchs
  • 6.1 Aeolidioidea
  • 6.2 Arminoidea
  • 6.3 Doridoxoidea
  • 6.4 Fionoidea
  • 6.5 Flabellinoidea
  • 6.6 Tritonioidea
  • 6.7 Unassigned families

• 7 Nematocysts and zooxanthellae

• 8 Conclusions

• 9 Conflicts of interest

• Acknowledgements

• References

---

Covering: up to the end of February 2017

Nudibranchs have attracted the attention of natural product researchers due to the potential for discovery of bioactive metabolites, in conjunction with the interesting predator-prey chemical ecological interactions that are present. This review covers the literature published on natural products isolated from nudibranchs up to February 2017 with species arranged taxonomically. Selected examples of metabolites obtained from nudibranchs across the full range of taxa are discussed, including their origins (dietary or biosynthetic) if known and biological activity.

---

Lewis J. Dean

Lewis Dean obtained his BSc(Hons) at the University of Waikato in 2016, completing his research dissertation on the natural products of New Zealand nudibranchs. A past winner of a prestigious University of Waikato Sir Edmund Hillary Scholarship, he was recently awarded a University of Waikato Doctoral Scholarship and is currently undertaking a PhD under the chief supervision of Michèle Prinsep, focusing on the isolation and characterisation of bioactive metabolites from New Zealand marine algae.

Michèle R. Prinsep

Michèle Prinsep received her BSc(Hons) and PhD degrees from the University of Canterbury, where she studied the isolation and structural elucidation of biologically active secondary metabolites from sponges and bryozoans under the supervision of Professors Blunt and Munro. She undertook postdoctoral research on cyanobacteria with Richard Moore at the University of Hawaii before returning to New Zealand to take up a lectureship at the University of Waikato, where she is currently an Associate Professor.

---

1 Introduction

Nudibranchs, often called sea slugs or more poetically, “butterflies of the sea”,¹ are a diverse group of marine gastropod molluscs, representing over 4700 known species.² After shedding their shells as larvae, nudibranchs thereafter remain shell-less; indeed the name nudibranch literally translates to “naked gill”, a reference to the exposed cerata on the backs of many species.³ Whilst essentially blind, nudibranchs perceive their environment through chemosensory interactions with two specialised rhinophores on their heads.^3,4 These carnivores are important consumers in benthic communities, feeding mostly upon sessile organisms, (sponges, cnidarians, tunicates and bryozoans), although some species hunt other nudibranchs. Both specialist and generalist feeders are known and can be found in practically all oceans.³

Having lost the physical protection of a shell, nudibranchs often utilise chemical defences to deter predators.³ Given that their typical prey species are some of the most prolific producers of natural products,⁵ it is perhaps unsurprising to find that many nudibranchs are known to sequester these defences.⁶ The vibrant colours associated with many nudibranch species are often correlated with those of their prey,⁶ however, de novo production of defences is also known.⁷ Nudibranchs have thus attracted many natural product researchers due to the potential for discovery of bioactive metabolites, in conjunction with the chemical ecological interactions between predator and prey that exist for many species.

2 Taxonomy

Broadly speaking, nudibranchs can be separated into two distinct groups based on their general morphology (Fig. 1)³ and digestive glands; dorids and aeolids.⁷ Dorid nudibranchs (clade: Euctenidiacea) have an intact digestive gland⁷ and are distinguished by a feather-like plume of gills on their dorsal side, circling the anus.³ They also commonly feature discrete pockets, bumps and other distortions, on their skin known as mantle dermal formations (MDFs), in which bioactive defence chemicals are typically stored.^3,8 Aeolid nudibranchs (clade: Cladobranchia) have a branched digestive gland⁷ and lack gills.³ They are characterised by the presence of dorsal projections known as cerata, which function in place of gills by facilitating gas exchange through the epidermis.³ In many aeolid species, the digestive tract also extends into the cerata and the tips often contain cnidosacs; stinging cells absorbed from prey species that are used for the nudibranch's own defence.³

Fig. 1 General morphology of a dorid nudibranch (Alloiodoris lanuginata) (left), highlighting the dorsal gill plumage and an aeolid nudibranch (Jason mirabilis) with characteristic cerata (right). Note the presence of rhinophores on both. Photographs courtesy of Tracey Bates. (After Picton and Morrow, 2006).³

Strictly speaking, a branched digestive gland is indicative of a non-dorid nudibranch, known as a cladobranch⁷ and true aeolids are only those belonging to the parvorder Aeolidida (Fig. 2), the only nudibranchs to possess cnidosacs.^7,9 However, many of the non-dorid nudibranchs are said to be aeolid-like as they possess cerata, or cerata-like projections, instead of gills.³ Others do possess gills, often tucked between the mantle and foot, but they bear little resemblance to the dorsal plumes of dorids.³

Fig. 2 Phylogenetic relationships between the taxonomic divisions currently recognised for Nudibranchia, and that are used in this review. Branches are terminated with superfamilies (underlined), whilst the roots represent taxonomic divisions (as used in WoRMS¹³).

The classification of nudibranchs is far from settled. There have been a number of taxonomic revisions in the past,^10–12 the one presented here being that currently used as in WoRMs.¹³ A clear distinction between dorid and non-dorid nudibranchs is well established, although a number of cladobranch families have yet to be assigned to an appropriate superfamily.¹³

Molecular studies are offering new insights into phylogenetic relationships. A relatively recent report analysing RNA sequence data, attempted to reconcile the groupings within Cladobranchia.¹⁴ Whilst many of the relationships highlighted in past classifications¹² are retained, some new clades have been suggested (although unnamed), incorporating some of the previously unassigned families. However, this analysis is by no means conclusive since only 10 of the 32 families currently recognised were investigated.¹⁴ A similar study analysed the mitochondrial and nuclear genes of the Doridoxa genus (the only genus in the current superfamily Doridoxoidea).¹⁵

The changes proposed by the above studies serve to highlight the ever evolving nature of taxonomy. This can lead to confusion for anyone attempting to navigate the literature covering nudibranch natural products, as invariably one will come across outdated phylogenetic relationships as well as old or repurposed taxonomic names. As an example, Doridacea is now recognised as an Infraorder within Euctenidiacea representing a subdivision of dorid nudibranchs,¹³ whereas historically it was used as a suborder covering all dorids.^3,7 Some species may also have been renamed and others folded into one another. This confusion has in some, thankfully rare, cases extended to a non-nudibranch sea slug (Pleurobranchaea meckeli) being reported as such.¹⁶

3 The origin of nudibranch natural products

Given that both sequestration and de novo synthesis of natural products are known in nudibranchs, it is rarely immediately clear if an isolated metabolite is of dietary origin. Feeding experiments with isotopically labelled precursors, are the only accepted procedures to prove biosynthesis of a metabolite.¹⁷ Unfortunately, the low uptake of feedstock by many marine organisms often forces the use of radioactive isotopes.¹⁷ Advances in nuclear magnetic resonance (NMR) spectroscopy have improved sensitivity and thus stable isotope experiments are increasingly becoming viable options, yet the cost of precursors, radioactive or stable, combined with the costs of very sensitive NMR instruments has inevitably restricted the use of feeding experiments in nudibranch studies.

There are many cases in the literature where de novo synthesis has been assumed, despite no feeding experiments being conducted. As the digestive glands of all nudibranchs are the source of nutrient distribution throughout the organism, it stands to reason that if a natural product is found therein, it is likely of dietary origin.⁷ Conversely if a metabolite is only present in the skin or mantle, and not in the viscera of a nudibranch, then it is probable that the compound is either biosynthesised in full or secondarily modified.⁷ Likewise, the absence or presence of the same, or structurally related, metabolite(s) in possible food species are also used to argue for biosynthesis or sequestration respectively.¹⁸ It has been further reasoned that any species of nudibranch that synthesises its own metabolites should consistently possess all metabolites, across all geographic collections.¹⁸ Species known to show variation in their natural products, especially from differing locations, may be indicative of metabolite sequestration.¹⁸ In addition, as one of the evolutionary advantages of de novo synthesis is to liberate a nudibranch from a singular food source, specialist feeders may be more likely to sequester metabolites.^7,18

4 Scope of review

There have been a number of excellent reviews of nudibranch chemistry over the years, including reviews on chemical ecology,^19–21 chemical defence^7,22,23 and bioactive metabolites.^24,25 Many focus on specific groups of metabolites such as diterpenes,²⁶ terpenoids,²⁷ cyanide and isothiocyanates²⁸ and isocyanides²⁹ (but all also include other marine invertebrates). A number of these reviews are regional in their focus, dealing with nudibranchs and other molluscs found in New Zealand,^30,31 Australia,^31,32 North America,³³ Japan,³⁴ Africa,³⁵ South America³⁵ and Antarctica.³⁵

The annual review of marine natural products that appears in Natural Product Reports has a section on molluscs^36a-f but given the wide scope of this review, there is no capacity to discuss nudibranchs in any detail or in a comprehensive manner over a considerable time period. There have only been three relatively recent reviews that deal with nudibranch chemistry, but two of these are not specific to nudibranchs (focus respectively on the terpene chemistry of marine molluscs³⁷ and marine molluscs as a source of antiviral drugs³⁸) and the third is limited in scope (deals with lipids and fatty acids of nudibranchs but only eight species were examined).³⁹

This current review focusses on the chemistry and chemical ecology of nudibranchs. It is arranged taxonomically and species are ordered as per their superfamily in the World Register of Marine Species (WoRMS)¹³ to highlight both the different and similar defensive strategies adopted by the various clades. Species are referred to and ordered by the name used in the original publication but where a taxonomic revision has taken place, the correct name is given in Table 1 and noted in brackets in the text. In each section, a selection of representative, significant or otherwise interesting, nudibranch natural products are discussed, including their origins (dietary or biosynthetic) if known and biological activity.

Table 1 Current classification (WoRMS) of nudibranchs covered in this review

Reported					Reclassification or (correction)
Superfamily	Family	Subfamily	Genus	Species
Dorids
Bathydoridoidea	Bathydorididae		Bathydoris	hodgsoni
Doridoidea	Actinocyclidae		Actinocyclus	papillatus
	Cadlinidae		Aldisa	andersoni
				sanguinea cooperi	Aldisa cooperi ^359
				smaragdina
			Cadlina	laevis
				luteomarginata
				pellucida
	Chromodorididae		Ardeadoris	egretta
			Casella	atromarginata	Doriprismatica atromarginata ^360
			Ceratosoma	amoena	Ceratosoma amoenum ^361
				brevicaudatum
				gracillimum
				trilobatum
			Chromodoris	albonotata
				albopunctata	Goniobranchus albopunctatus ^362
				annulata	Goniobranchus annulatus ^363
				cavae	Goniobranchus cavae ^364
				elisabethina
				epicura
				funerea	Chromodoris lineolata ^365
				geminus	Goniobranchus geminus ^366
				geometrica	Goniobranchus geometricus ^367
				gleniei	Goniobranchus gleniei ^368
				hamiltoni
				inopinata
				inornata	Chromodoris aspersa ^369
				kuniei	Goniobranchus kuniei ^370
				lochi
				luteorosea	Felimida luteorosea ^371
				macfarlandi	Felimida macfarlandi ^372
				maridadilus	Hypselodoris maridadilus ^373
				marislae	Felimida marislae ^374
				michaeli
				norrisi	Felimida norrisi ^375
				obsoleta	Goniobranchus obsoletus ^376
				petechialis	Goniobranchus petechialis ^377
				quadricolor
				reticulata	Goniobranchus reticulatus ^378
				sedna	Doriprismatica sedna ^379
				sinensis	Goniobranchus sinensis ^380
				tinctoria	Goniobranchus tinctorius ^381
				willani
				youngbleuthi	Glossodoris rufomarginata ^382
			Felimida	grahami	(Felimare grahami)^383
			Glossodoris	atromarginata	Doriprismatica atromarginata ^360
				averni	Ardeadoris averni ^384
				cincta
				dalli	Felimida dalli ^385
				hikuerensis
				pallida
				quadricolor	Chromodoris quadricolor ^386
				rufomarginata
				sedna	Doroprismatica sedna ^379
				tricolor	Felimare tricolor ^387
				valenciennesi	Felimare picta ^388
				vespa
			Goniobranchus	albonares
				collingwoodi
				daphne
				splendidus
				verrieri
			Hypselodoris	agassizii	Felimare agassizii ^389
				californiensis	Felimare californiensis ^390
		cantabrica		Felimare cantabrica ^391
daniellae		Thorunna daniellae ^392
fontandraui		Felimare fontandraui ^393
ghiselini		Felimare ghiselini ^394
godeffroyana		Risbecia godeffroyana ^395
infucata
jacksoni
lajensis		Felimare lajensis ^396
orsini(i)	Felimare orsinii ^397
porterae	Felimare porterae ^398
tricolor	Felimare tricolor ^387
villafranca	Felimare villafranca ^399
webbi	Felimare picta ^400
zebra	Felimare zebra ^401
Miamira		magnifica
		miamirana
Risbecia	tryoni	Hypselodoris tryoni ^402
Tyrinna	nobilis	Tyrinna delicata ^403
Discodorididae	Anisodoris	nobilis	Peltodoris nobilis ^404
		fontainei	Doris fontainii ^405
	Asteronotus	cespitosus
	Diaulula	sandiegensis
	Halgerda	aurantiomaculata
		gunnessi
		rubicunda	Sclerodoris rubicunda ^406
		stricklandi
		theobroma
		willeyi
	Jorunna	funebris
	Peltodoris	atromaculata
	Platydoris	sp.
	Sclerodoris	tanya
Dorididae	Archidoris	carvi	Doris fontainii ^405
	Neodoris	carvi	Doris fontainii ^405
	Archidoris	montereyensis	Doris montereyensis ^407
		odhneri	Doris odhneri ^408
		pseudoargus	Doris pseudoargus ^409
		tuberculata	Doris pseudoargus ^409
	Austrodoris	kerguelenensis	Doris kerguelenensis ^410
	Doris	verrucosa
Onchidoridoidea	Goniodorididae	Hopkinsia	rosacea	Okenia rosacea ^411
		Okenia	zoobotryon
	Onchidorididae	Acanthodoris	nanaimoensis
		Adalaria	loveni
Phyllidioidea	Dendrodoridae	Dendrodoris	arborescens
			carbunculosa
			denisoni
			fulva
			grandiflora
			krebsii
			limbata
			nigra
			tuberculosa
		Doriopsilla	albopunctata
			areolata
			janaina
			pelseneeri
	Phyllidiidae	Phyllidia	bourgini	Phyllidiella rosans ^412
			coelestis
			ocellata
			pulitzeri
			pustulosa	Phyllidiella pustulosa ^413
			varicosa
		Phyllidiella	rosans
		Phyllidiopsis	krempfi
		Reticulidia	fungia
Polyceroidea	Aegiridae	Notodoris	citrina
			gardineri
	Hexabranchidae	Hexabranchus	sanguineus
	Polyceridae	Nembrothinae	Nembrotha		kubaryana
			Roboastra		tigris
			Tambja		abdere
					ceutae
					eliora
					stegosauriformis
		Polycerinae	Polycera		tricolor
		Triophinae	Limacia		clavigera
			Triopha	carpenteri	Triopha catalinae ^414
				catalinae
Cladobranchs
Aeolidioidea	Aeolidiidae		Aeolidia	papillosa
			Aeolidiella	stephanieae	Berghia stephanieae ^415
			Spurilla	sp.
	Facelinidae		Cratena	peregrina
			Hermissenda	crassicornis
			Phidiana	militaris
			Phyllodesmium	guamensis	(Phyllodesmium guamense)^416
				lizardensis	(Phyllodesmium lizardense)^417
				longicirra	(Phyllodesmium longicirrum)^418
				magnum
			Pteraeolidia
Arminoidea	Arminidae		Armina	maculata
			Dermatobranchus	ornatus
	Doridomorphidae		Doridomorpha	gardineri
Doridoxoidea
Fionoidea	Fionidae		Phestilla	melanobrachia	Tenellia melanobrachia ^419
				sibogae	Tenellia sibogae ^420
Flabellinoidea	Flabellinidae		Flabellina	affinis
			Flabellinopsis	iodinea	Flabellina iodinea ^421
Tritonioidea	Tethydidae		Melibe	leonina
			Tethys	fimbria
	Tritoniidae		Marionia	limceana
			Tritonia	diomedea	Tritonia tetraquetra ^422
				hamnerorum
	Tritoniidae		Tritoniella	belli
			Tritoniopsis	elegans
			Tochuina	tetraquetra	Tochuina gigantea ^423
Unassigned
Families	Charcotiidae		Charcotia	granulosa
			Leminda	millecra
	Dotidae		Doto	antarctica
				carinova
				pinnatifida
	Pinufidae		Pinufius	rebus
	Proctonotidae		Janolus	cristatus
				novozelandicus

---

5 Dorid nudibranchs

5.1 Bathydoridoidea

5.1.1 Bathydorididae. Bathydoridoidea is a small superfamily of deep water nudibranchs containing only the family Bathydorididae, comprised of one genus, Bathydoris. It has received little attention from natural product chemists, presumably due to difficulties in collecting specimens. A single investigation of one species has been reported to date. Hodgsonal (1), a novel, drimane sesquiterpene was isolated from the mantle of Bathydoris hodgsoni⁴⁰ and was the first known 2-substituted drimane sesquiterpene of marine origin.^40,41 The distribution of 1 among individual B. hodgsoni specimens collected from various sites was found to be reasonably consistent and it was only detected on the extremities of the nudibranch and not in the viscera. Given that the stomach contents of B. hodgsoni indicated that this species is a generalist and opportunistic feeder, it was suspected that 1 is de novo synthesised.⁴¹ In experiments, 1 deterred feeding in a potential starfish predator (Odontaster validus) at natural concentrations, highlighting its probable use as a defence allomone.⁴¹

5.2 Doridoidea

Comprising five families, Doridoidea is by far the most studied nudibranch superfamily in terms of species, genera and metabolites reported. Searching within the MarinLit database⁴² for natural products investigations into nudibranchs (Taxonomy filter: Phylum-Mollusca; Article free text filter: nudibranch) reveals 152 records (as of 28 February 2017). Of these, 80 concern species of Doridoidea (56%).

5.2.1 Actinocyclidae. Actinocyclidae is a small family comprising two genera (Actinocyclus and Haliaxa) and only one study has been conducted on a species in this family. An isonitrile lipid, actisonitrile (2) was obtained from the mantle of a South China Sea specimen of Actinocyclus papillatus as the major component. Total synthesis was achieved and both enantiomers exhibited moderate cytotoxicity to a mammalian cell line, which, along with the selective distribution of 2, implies a likely defensive function for the metabolite.⁴³

5.2.2 Cadlinidae. The Cadlinidae family comprises two genera (Aldisa and Cadlina) and the chemistry of representatives of both has been studied. An Indian collection of Aldisa andersoni was the source of some members of the phorbazole family of metabolites,⁴⁴ including the new 9-chloro-phorbazole D (3) and N1-methyl-phorbazole A (4), which co-occurred with known phorbazoles A, B and D, previously obtained from the sponge Phorbas aff. clathrata.⁴⁵ Although found mainly on the exterior of A. andersoni, the compounds were also present in the digestive organs of the mollusc, reinforcing the high probability that these compounds were sequestered from a Phorbas sponge, although none were observed near the collection site.⁴⁴ Phorbazole A, 3 and 4 deterred feeding by the shrimp Palaemon elegans and both 3 and 4 exhibited cytostatic effects against several human tumour cell lines (HTCLs).⁴⁴

Two steroidal acids, 3-oxo-chol-4-ene-24-oic acid (5) and its unsaturated analogue (6), were isolated from Aldisa sanguinea cooperi⁴⁶ (Aldisa cooperi³⁵⁹). Cholestenone (7), an alkyl derivative of 5 and 6 was also present in the extract. A. cooperi was consistently found feeding upon the sponge Anthoarcuata graceae, which, whilst lacking 5 and 6, was found to contain 7 as one of its major metabolites. Cholestenone, 7, proved to be inactive in feeding experiments whereas 5 was an effective inhibitor of feeding behaviour (common goldfish). This indicated that A. cooperi was obtaining an inactive metabolite from its diet and modifying it to provide a defence against predators.⁴⁶ It has been suggested that another steroidal metabolite, 24-norchol-4-ene-3,22-dione (8), isolated from the related nudibranch A. smaragdina may also be secondarily modified from 7 (ref. 47) or alternatively, result from the β-oxidation and subsequent decarboxylation of 5. However, these suggestions were tentative since no steroidal precursors were noted in the observed prey species Phorbas fictitius.⁴⁷

A nudibranch that has attracted significant interest is Cadlina luteomarginata, as it is one of only two species of nudibranchs known to both sequester prey metabolites and biosynthesise its own natural products⁴⁸ (the other, Dendrodoris grandioflora, will be discussed below). To date, 38 terpenoid metabolites, with 22 carbon skeletons, representing monoterpenes,⁴⁹ sesquiterpenes,^49–51 diterpenes,^52,53 sesterterpenes⁵⁴ and degraded sesterterpenoids^51,55 and diterpenoids^53,56 have been isolated from C. luteomarginata.⁵⁷ Examples of the sequestered metabolites are glaciolide (9)⁵⁶ and cadlinolide A (10)⁵³ from the sponge Aplysilla glacialis, and ansellone A (11)⁵⁴ from a prey sponge of the Phorbas genus. Their ecological roles are not well defined,⁵⁷ but 11 has been shown to be moderately activating towards cellular processes that use the cyclic adenosine monophosphate (cAMP) signal pathway.⁵⁴

Terpenoid biosynthesis in C. luteomarginata was suspected when all previously analysed individuals from a British Columbian (Canadian) study consistently possessed the same three terpenoids.⁵⁸ The de novo production of these terpenes; albicanyl acetate (12), cadlinaldehyde (13) and luteone (14) was subsequently proven by stable isotope labelling experiments.⁵⁸ Antifeeding activity of 12 was detected, and it was found at high concentration in the mantle and mucus, indicating its role as a deterrent.⁵¹ A derivative of 12, 1α,2α-diacetoxyalbicanyl acetate (15), was isolated from the egg masses of C. luteomarginata but not detected in the nudibranch itself.⁵⁷ Whilst no feeding assays of 15 were conducted, its similarity to 12 indicated that it may act to protect the egg masses.⁵⁸ Likewise de novo synthesis may be expected for 15, despite it not being detected to date in C. luteomarginata nudibranchs.

Initially, there was speculation that C. luteomarginata may be divided into subspecies based on geographic distribution, as a Californian population was originally thought not to contain any of the biosynthesised compounds 12–14.^48,58 However, reexamination of this population utilising gas chromatography (GC) indicated that 12–14 were present but at low concentrations.⁴⁸ It was found that broadly speaking, individuals of C. luteomarginata, from British Columbia, could be separated into two groups. The first group possessed 12–14 at high concentration, with other terpenoids that could be detected by GC at much lower concentration, whilst the second group contained 12–14 at concentrations below those of other GC detectable terpenoids. This variance both between and within geographic populations suggested that C. luteomarginata was able to downregulate biosynthesis if sufficient dietary terpenoids could be sequestered.⁴⁸

A number of terpenoids were obtained from C. pellucida and C. laevis, including the furanosesquiterpene laevidiene (16) from the latter. The structures indicated a dietary origin from sponge prey, although only C. pellucida was observed feeding upon a sponge (Spongia agaricina).⁵⁹

5.2.3 Chromodorididae. Chromodorididae is a large family with 17 accepted genera, 11 of which have been investigated chemically. The majority of natural product studies of nudibranchs have been conducted on this family, especially on the genus Chromodoris. There has been one reported study of a nudibranch from the genus Ardeadoris. An Australian collection of A. egretta yielded a new diterpene (17) but its bioactivity was not tested.⁶⁰

Six furanoditerpenoids were obtained from Casella atromarginata (Doriprismatica atromarginata³⁶⁰) from Sri Lanka.⁶¹ Of these, two were known metabolites of an Australian Spongia sp.⁶² and two more were minor structural variants but two (18 and 19) were novel metabolites, with a more highly oxidised A-ring, containing a monoenolised α-diketone function.⁶¹ It seems likely that C. atromarginata obtained these metabolites from its sponge prey.⁶¹

Ceratosoma amoena (Ceratosoma amoenum³⁶¹) from New Zealand was observed on the red alga Hymenea variolosa.⁶³ The red algal metabolite allolaurinterol⁶⁴ (20) was isolated from both the nudibranch and alga, a surprising result since, as noted by the researchers, the nudibranch dietary system is not capable of breaking down algal tissue. As an alternative explanation for this result, it was suggested that C. amoenum could be preying on an organism (or more likely its egg masses) that does sequester this compound from the alga.⁶³

Nine new spongiane diterpenes were isolated from a South Australian nudibranch tentatively identified as Ceratosoma brevicaudatum⁶⁵ but later identified as Chromodoris epicura⁶⁶ so perhaps it is unsurprising that these do not seem to be typical of the Cerastoma genus. A thiosesquiterpene (21) was isolated from the Australian nudibranch C. brevicaudatum but believed to be sequestered from a Dysidea sponge.⁶⁷ The ecological role of 21 has not been studied, but it represents a relatively rare example of a sulphur containing sesquiterpene from the marine environment.⁶⁷ Four “typical” sponge furanosesquiterpenoids, pallescensin-B,⁶⁸ (−)-furodysinin,⁶⁹ dehydroherbadysidolide⁷⁰ and herbadysidolide⁷¹ were isolated from two species of Ceratosoma; C. trilobatum and C. gracillimum.⁶⁶ As previously reported,⁷² (−)-furodysinin, exhibited feeding deterrent and ichthyotoxic properties, and additionally, concentration of the compound in the dorsal horn glands of the animals, supports the proposed defensive role of the dorsal horns.⁶⁶

Chromodoris is the most examined of all nudibranch genera, with many reported studies,^{8,39,73–106} although many of the species in these studies originally classified as Chromodoris have been reclassified as Doriprismatica,⁷⁵Felimida,^{73,76,79,80,85,87,90,93}Glossodoris^79,101Goniobranchus^{39,86,88,89,95,98,102–104} or Hypselodoris.^74,106Chromodoris species feed exclusively on sponges and sequester a large number of terpenoid metabolites from their prey including sesquiterpenes,^{73,74,78,82,106} norditerpenes,⁸⁰ diterpenes^{8,76,81,85–90,93,95,96,98,99,102–104} and sesterterpenes,^{79,84,91,92,92,97,100} although macrolides^77,82,83,105 and bromophenols⁷⁸ have also been reported. In some instances, metabolites have been simultaneously isolated from sponge and nudibranch, highlighting that they are of dietary origin. For example, the macrolides laulimalide (22)^107,108 and isolaulimalide (23)¹⁰⁹ were isolated from sponges of the Hyattella genus and from their nudibranch predator C. lochi.⁸³

Latrunculin A, a sponge metabolite originally obtained from Latrunculia magnifica¹¹⁰ has been isolated from several Chromodoris species,^77,82,94,105 and has been shown to be selectively stored in the mantle rim for protective purposes.¹⁰⁵ In other cases, structural analogues of nudibranch metabolites have been detected in or isolated from, the sponge prey, indicating the likelihood of secondary modification of sponge metabolites. For example, the sesquiterpene marislin (24), obtained from C. marislae (Felimida marislae³⁷⁴), is related to the spongian metabolite pleraplysillin-2 (25), from Pleraplysilla spinifera, by a [3,3] sigmatropic rearrangement.⁷³ Similarly, deoxymanoalide (26) and deoxysecomanoalide (27), isolated from C. willani, are modified from manoalide (28) and secomanoalide (29) respectively by loss of an hydroxyl group. Both 28 and 29 were identified in the sponge prey of C. willani.¹⁰⁰ Some chlorinated homoditerpenes including hamiltonin A (30)⁹⁴ were isolated from C. hamiltoni along with latrunculins and a sesterterpene but in this case, the origin of the chlorinated compounds was not clear.⁹⁴ Where they have been assessed, virtually all of the metabolites obtained from Chromodoris species have displayed a moderate degree of bioactivity, indicating their role in limiting predation. Despite evidence suggesting secondary modification occurs, there have been no reports of de novo synthesis within Chromodoris to date.

A species, denoted by researchers as Felimida grahami (Felimare grahami,³⁸³) sequestered rearranged terpenoids from its sponge prey Darwinella cf. oxeata.¹¹¹ These included oxeatamide I (31), one of three new oxeatamides H–J¹¹¹ found in the sponge to add to those already known¹¹² and membranolide,¹¹³ previously isolated from the sponge Dendrilla membranosa¹¹³ and from nudibranchs Goniobranchus reticulatus¹⁰² and G. splendidus.¹¹⁴

As for Chromodoris, many of the species originally reported as Glossodoris have been reclassified, in this case as Ardeadoris,¹¹⁵Chromodoris,¹¹⁶Doriprismatica,^39,117–120Felimare¹²¹ and Felimida.¹²⁰ Although latrunculin B has been isolated from G. tricolor, this is now classified as Felimare³⁸⁷ and all other metabolites isolated from this genus are terpenoids and predominantly sponge derived.^{115,117–126} Spongian diterpenes have been obtained,^117,119 but most of the metabolites isolated are based on the sponge-derived scalarane sesterterpene, scalaradial (32),^{115,119,121–125} including norscalaranes¹¹⁹ and homoscalaranes.¹²⁰ A number of the scalarane metabolites obtained were 12-keto derivatives such as 33 ¹²⁴ and since in every case, they were only obtained from the nudibranch and not from the sponge prey, it is considered highly likely that biotransformation of the sponge scalarane occurs in the nudibranch,¹²⁴ most likely through enzymatic oxidation.¹¹⁵

As noted above, taxonomic revisions have led many species that were formerly Chromodoris to be reclassified as Goniobranchus. From species originally reported as Goniobranchus, a range of diterpenes and norditerpenes have been obtained and all studies thus far have been of Australian specimens.^{60,114,126–128} In most cases, known sponge metabolites were co-isolated with the novel metabolites reported and compounds were sometimes isolated from both the nudibranch and its prey sponge. 12-Acetoxy dendrillolide (34) for example was obtained from both G. albonares and the crimson sponge on which it was feeding.¹²⁶ The activity of isolated metabolites has been quite varied. For example, of the six new oxygenated norditerpene/bisnorditerpene metabolites isolated from G. splendidus, gracilins M–Q displayed significant potency against the HeLa S3 cell line, aplytandiene-3 was essentially inactive and the rearranged diterpene daphnelactone (35) reported from G. daphne in the same study was not tested.¹¹⁴ Another study of G. splendidus yielded some oxygenated diterpenes but these were not tested for activity.⁶⁰G. verrieri yielded seven new norditerpenes and diterpenes, all with highly rearranged carbon skeletons, such as the cyclopropyl-containing verrielactone (36) and a biosynthetic pathway to these metabolites from spongialactone was proposed.¹²⁷ All six new spongian-16-one diterpenes found localised in the mantle of G. collingwoodi (an example being 37) were inactive to a range of HTCLs but preliminary testing of whole body extracts indicated some antifeedant activity against Palaemon serenus (rock pool shrimp).¹²⁸

The majority of species originally reported as Hypselodoris have been reclassified as Felimare^{8,72,129–138} or in one case, Risbecia¹⁰⁶ but the genus sequesters terpenes,^{8,72,98,106,129–139} predominantly furanosesquiterpenes,^{8,72,98,106,134–139} from various species of sponge, of which longifolin¹³⁸ (38) is typical. Longifolin (38) is sequestered from Dysidea sponges^{106,130,136,137} and it and other antifeedant compounds are present in the MDFs of many Hypselodoris species^8,72,131 and released when the nudibranch is molested. There is one reported isolation of a sesterterpene from this genus.¹³³H. orsini(i) (Felimare orsinii)³⁹⁷ feeds on the sponge Cacospongia mollior but contains different metabolites to its prey. It is thought that the nudibranch converts the sponge sesterterpenoid scalaradial, into deoxoscalarin, then, a second chemical transformation produces the sesterterpenoid, 6-keto-deoxoscalarin which is also stored in MDFs.¹³³

A number of oxy-polybrominated diphenyl ethers (O–PBDEs) have been isolated from the nudibranchs Miamira magnifica and M. miamirana from Australia, of which 39 from M. magnifica is a typical example.¹⁴⁰ These compounds are sequestered from sponges and although such compounds have been isolated from nudibranchs previously,^78,141 they were found in the gut tissue, whereas in the Miamira specimens, they were additionally located in the mantle and dorsal horn tissues, so likely play a role in chemical defence.¹⁴⁰

As part of a study of Chromodorid species,⁸Risbecia tryoni (Hypselodoris tryoni⁴⁰²) yielded the sponge metabolite (−)-furodysinin¹⁴² and the lipid and fatty acid content has also been examined.³⁹ There has only been one chemical study of the Tyrinna genus. Tyrinnal (40), a seco-11,12-spongiane was obtained from the mantle and MDFs of Patagonian Tyrinna nobilis (Tyrinna delicata⁴⁰³) along with three known furanosesquiterpenes.¹⁴³

5.2.4 Discodorididae. The Discodorididae are a large family of nudibranchs consisting of 30 genera. Only eight of these have been chemically examined (and some of these examples have been reclassified to other genera within the family,^144,145 or in one case, to a genus in the Dorididae family¹⁴⁶) but they have yielded metabolites from a range of classes.

A number of diterpenoid diacylglycerols have been isolated from this family, including the anisodorins 1–5, which were obtained from the skin of the Patagonian species Anisodoris fontainei (Doris fontainii⁴⁰⁵) along with two further analogues.¹⁴⁶ These last two compounds were previously reported from the same species¹⁴⁷ (but erroneously classified as Archidoirs carvi¹⁴⁶) and are diastereoisomers of two further diacylglycerols obtained from Archidoris tuberculata (Doris pseudoargus⁴⁰⁹), collected along the northern Spanish coast.¹⁴⁷ Surprisingly, the isomers from each source display opposite absolute stereochemistry in their terpenoid moiety. The absolute stereochemistry of anisodorin 5, 41 was established through synthesis of its enantiomer¹⁴⁸ and 41 itself has also been synthesised.¹⁴⁹Anisodoris nobilis (Peltodoris nobilis⁴⁰⁴) was the source of doridosine,¹⁴⁴ (1-methylguanosine), a hypotensive compound¹⁵⁰ that was also obtained from the Australian sponge Tedania digitata.¹⁵¹ A degraded sesquiterpenoid was obtained from A. nobilis as the odoriferous principle.¹⁵²

Asteronotus cespitosus specimens collected from Australia and the Philippines contained a range of halogenated metabolites and two sesquiterpenes.¹⁴¹ One of these was a novel chlorinated pyrrolidone (42), while all of the other metabolites had been previously isolated from Dysidea herbacea.¹⁴¹ Pyrrolidone (42) has a similar structure to dysideapyrrolidone,¹⁵³ a sponge metabolite.

Isoguanosine, a compound previously isolated as a constituent of the croton bean¹⁵⁴ was obtained from Diaulula sandiegensis.¹⁵⁵ A Californian population of D. sandiegensis contained a series of nine chlorinated acetylenes, of which 43 is a typical example.¹⁵⁶ Over twenty years later, six of the metabolites were isolated from the sponge Haliclona lunisimilis which was collected in the same area as the nudibranchs were originally.¹⁵⁷ The two major metabolites found in D. sandiegensis (both ketones) were not present in the sponge, although the corresponding alcohols were, suggesting modification of the alcohols by the nudibranch.¹⁵⁷ Steroidal metabolites, diaulusterols A (44) and B (45), have been isolated from a British Columbian collection of D. sandiegensis.¹⁵⁸ Stable isotope experiments indicated that the polyketide chain of 44 was biosynthesised by the nudibranch itself, whereas there was no evidence of uptake in the steroidal nucleus.¹⁵⁹ Radiolabelled incorporation also failed to prove biosynthesis of the steroid nucleus.¹⁵⁹ This suggested that as for Aldisa cooperi,⁴⁶ a precursor steroid was sequestered and subsequently modified, although a prey species has yet to be identified.^158,159

Five species of the Halgerda genus collected from various locations around Australia and from Okinawa, Japan were examined. Of these, H. aurantiomaculata yielded the new tryptophan derivative halgerdamine (46), in addition to four known nitrogenous compounds, while H. gunnessi yielded mixtures of acylated tetrasaccharides.¹⁴⁵ No secondary metabolites were detected in H. theobroma, H. rubicunda (Sclerodoris rubicunda⁴⁰⁶) or H. willeyi, leading the authors to suggest that these nudibranchs may have evolved to exploit a wider range of food including nontoxic sponges rather than sequestering or synthesising protective compounds.¹⁴⁵ Organic extracts of an Indian specimen of H. stricklandi displayed modest activity to Staphylococcus aureus and no activity against a range of other bacterial and fungal species.¹⁶⁰

Jorunna funebris is another nudibranch to yield non terpenoid natural products. Studies of the nudibranch and its prey, sponges of the genus Xestospongia, indicate that J. funebris sequesters a range of isoquinolinequinones^160–163 and bistetrahydroisoquinolines^162,164,165 from the sponge. Some of the isoquinolinequinones have been synthesised¹⁶⁶ and the metabolites possess a range of biological activities, including antibacterial activity,^161,164 cytotoxicity to HTCLs,^162,164,165 and NF–κB inhibitory activity.¹⁶² Jorumycin (47) was not found in the sponge but was isolated from the mucus and mantle of an Indian collection of J. funebris. It exhibited very potent cytotoxicity towards various tumour types but was only available in small quantities due to instability of the carbinolamine moiety.¹⁶⁴ Pretreatment of the nudibranch tissues with potassium cyanide, resulted in isolation of jorunnamycins A–C (C pictured as 48), more stable compounds which retained high cytotoxicity to human tumour cell lines.¹⁶⁵ Zalypsis®, a synthetic compound derived from jorumycin,¹⁶⁷ is currently in Phase II clinical trials for treatment of endometrial and cervical cancer.¹⁶⁸

Several studies have chemically linked the nudibranch Peltodoris atromaculata to its prey sponges Petrosia ficiformis^169,170 and Haliclona fulva,¹⁷¹ indicating sequestration of sponge metabolites by P. atromaculata, including sterols,¹⁶⁹ the polyacetylene alcohols, petroformynes¹⁷⁰ and fulvinol-related polyacetylenes.¹⁷⁰ One of these last compounds was moderately cytotoxic to the SKMEL-28 melanoma cell line.¹⁷¹ The lipid and fatty acid content of the related Platydoris sp. were examined as part of a study of eight nudibranch species.³⁹

The sesquiterpene glyceride esters, tanyolides A and B were isolated from Sclerodoris tanya from Southern California. Located mainly in the mantle of the nudibranch, both deterred feeding by fish predators and were also synthesised.¹⁷² Feeding experiments utilising [2-¹³C]mevalonolactone indicated that Sclerodoris tanya is capable of de novo biosynthesis of the terpenoid fragment of tanyolide B.¹⁷³

5.2.5 Dorididae. The Dorididae family consists of six genera. All Dorididae species originally classified as Archidoris and Austrodoris have been reclassified as Doris^{405,407–410} so technically only this genus has been chemically examined, but there have been quite a number of reports published under the original genera names.^{147,152,173–180} Given the reclassification, it is unsurprising that terpenoid glyceryl esters have been reported from all three genera and seem to serve as “chemical markers” of the family.¹⁷⁴ Species originally classified as Archidoris have yielded sesquiterpenoic^152,175,176 and diterpenoic acid^{147,152,176–178,180} glycerides, in addition to a glyceryl ether.¹⁵² Archidorin (49)¹⁷⁸ is a typical example. Many of these compounds were isolated from the skin or mantle of the nudibranchs and were icthyotoxic,^{147,152,177,178,180} and in one study, were additionally shown to be passed from the adults into the egg masses for protection.¹⁸⁰ Diastereoisomeric variants of some of the diterpenoic acid glycerides were obtained from geographically distinct populations of Archidoris nudibranchs¹⁴⁷ and some of these compounds have been synthesised.^174,179 Evidence from biosynthetic feeding experiments indicate that the nudibranchs likely synthesise these molecules de novo.¹⁷³

The Antarctic nudibranch Austrodoris kerguelenensis (Doris kerguelenensis⁴¹⁰) has yielded a number of diterpene glycerides with a variety of skeletons, even within one population,¹⁸¹ including ent-labdane,^182,183 labdane,¹⁸⁴ halimane,^184,185 clerodane^184,186,187 and isocopalane¹⁸⁸ frameworks.^186,189 Some norsesquiterpenes¹⁹⁰ have also been isolated from this well-studied species. Of the compounds isolated, some of the diterpene diacylglycerides had feeding deterrent activity against the predatory seastar Odontaster validus,¹⁹¹ some of the clerodane (palmadorins A and B) and labdane (palmadorins M–O) diterpenes were growth inhibitory to human erythroleukamia (HEL) cells and palmadorin M (50) was shown to be apoptotic to these cells.¹⁸⁴ Not surprisingly, as for Archidoris, it is thought likely that the above compounds are biosynthesised de novo by the nudibranch.^191,192

The verrucosins are also diterpenoid acid glycerides obtained from Doris verrucosa,^193,194 and in addition to icthyotoxicity, verrucosins A and B (51 and 52) were shown to be potent activators of protein kinase C and to promote tentacle regeneration in the freshwater hydrozoan Hydra vulgaris.¹⁹⁵ Biosynthetic feeding experiments with ¹⁴C labelled mevalonic acid and glycerol resulted in poor incorporation into the verrucosins so de novo biosynthesis was not proven but good incorporation into sterols did establish that the mevalonate pathway was operative in the nudibranch.¹⁹⁶ A xylosyl thioether, 9-[5′-deoxy-5′(methylthio)-β-D-xylofuranosyl]adenine was also obtained from D. verrucosa^197–199 and biosynthetic studies indicated that it arises in D. verrucosa from isomerisation of endogenous 5′-deoxy-5′-methylthioadenosine (MTA).²⁰⁰

5.3 Onchidoridoidea

The Onchidoridoidea consists of five families but only representatives of the Onchidorididae and the Goniodorididiae have been examined for natural products.

5.3.1 Onchidorididae. The family Onchidorididae comprises five genera and has been little studied chemically. Only one species from each of two genera (Acanthodoris and Adalaria) has been examined. Acanthodoris nanaimoensis from British Columbia yielded three sesquiterpenoid aldehydes with novel skeletons, nanaimoal,²⁰¹ acanthodoral²⁰² and isoacanthodorane.²⁰² Biosynthetic studies involving injecting the animals with [1,2-¹³C₂]acetate, indicated that the compounds were biosynthesised de novo by the nudibranchs and a biosynthetic scheme was proposed.²⁰³ A degraded triterpenoid, lovenone (53) was isolated from skin extracts of a Norwegian collection of Adalaria loveni and displayed modest cytotoxicity to two HTCLs. A dietary origin was suspected, given the lack of any such metabolites in a closely related species.²⁰⁴

5.3.2 Goniodorididiae. The Goniodorididiae family comprises eight genera of which only one, Hopkinsia, has been examined for natural products. Hopkinsia rosacea (Okenia rosacea⁴¹¹) yielded a seemingly unique apocarotenoid, hopkinsiaxanthin which is similar to the carotenoid fucoxanthin found in marine plants.^205,206 The alkaloid 2,5,6-tribromo-N-methylgramine was obtained from the nudibranch Okenia zoobotryon and from its bryozoan prey, Zoobotryon verticillatum¹³⁷ (Amathia verticillata²⁰⁷).

5.4 Phyllidioidea

The Phyllidiodea superfamily comprises three families (Dendrodoridae, Mandeliidae and Phyllidiidae) and is very well studied, although no representatives of the Mandeliidae have been studied to date. All metabolites isolated thus far from this superfamily have been terpenes.

5.4.1 Dendrodoridae. Two of the three genera comprising the Dendrodoridae family (Dendrodoris and Doriopsilla) have been chemically examined. The discovery of de novo synthesis of the drimane sesquiterpene polygodial (54) in Dendrodoris limbata was the first proven example of biosynthesis in a nudibranch.²⁰⁸ The antifeeding properties of 54, in conjunction with its localisation solely within the mantle, indicated its defensive role.²⁰⁸ Surprisingly, 54 was found to be toxic to D. limbata if injected directly into the hepatopancreas,^209,210 presumably due to its reactivity with free amino groups.^208,210 It was speculated that a protected form of 54 must be produced in order to facilitate transport throughout the nudibranch. Olepupuane (55) was also found in D. limbata and was converted to 54 in the presence of aqueous acid, suggesting that 55 was the protected form.²¹⁰ A mixture of three fatty acid esterified sesquiterpenoids isolated from D. limbata may offer an alternative storage or handling method of the toxic 54 for the nudibranch.²¹¹

Biosynthesis of 54 and related compounds has been proven in Dendrodoris grandiflora through radiolabelling experiments.^212,213 Whilst 54 was extracted from the mantle, the fatty acid esterified sesquiterpenoids were found in the digestive glands along with a series of known spongian terpenes.²¹² Other studies have reported drimane sesquiterpenes from Dendrodoris carbunculosa,²¹⁴D. nigra, D. tuberculosa and D. krebsii.^215,216 The anatomical distribution and role that such terpenes may play has been investigated in Mediterranean Dendrodoris nudibranchs.²¹⁷ The presence of sequestered sponge metabolites might indicate a dual defence for Dendrodoris species but they were completely absent from the mantle, where one would expect to find defence allomones. Unlike Cadlina luteomarginata, in which it appears de novo synthesis is downregulated when sequestered metabolites are available,⁴⁸D. grandiflora still utilises its own natural products for defence.

Drimane biosynthesis has also been reported in Dendrodoris arborescens, in the form of 56, following radiolabelling studies and another drimane, 6β-acetoxypolygodial (57), was also isolated from the mantle of D. arborescens.²¹³ Feeding experiments with ¹⁴C- and ¹³C-labelled glucose on D. limbata and D. grandiflora indicated that glucose was incorporated into the terpenoid portion of drimanes via the standard mevalonate pathway.²¹⁸ A more recent study identified the presence of symbiotic bacteria in the outer tissues of D. nigra and it was suggested that they could be implicated in defence from predators.²¹⁹

In a New Zealand study, the sesquiterpene cinnamolide (58) was detected in all specimens of D. denisoni examined, regardless of geographic location or age, potentially indicating that it was biosynthesised by the nudibranch.⁶¹ Cinnamolide (58) has previously only been obtained from terrestrial plants.²²⁰

Studies of species of the related Doriopsilla genus including D. albopunctata^215,216D. janaina,²¹⁵D. pelseneeri²²¹ and D. areolata^{211,222–224} indicated that they also contained drimane sesquiterpenoids, as well as enantiomerically related sesquiterpenoids of the ent-pallescensin A skeleton.^221–224 Biosynthetic investigations of D. areolata^222,223 and D. pelseneeri²²⁵ indicated that they produce both drimane sesquiterpenoids and sesquiterpenoids with the ent-pallescensin A skeleton de novo and their presence in the mantle of D. areolata indicated that they too were involved in the chemical defence of the nudibranch.^222,223

5.4.2 Phyllidiidae. Representatives of four of the five genera comprising the Phyllidiidae family (Phyllidia, Phyllidiella, Phyllidiopsis and Reticulidia) have been chemically studied. Despite de novo synthesis being well documented among the superfamily Phyllidioidea, sequestering of metabolites still occurs, especially within the genus Phyllidia. A number of highly bioactive metabolites have been reported from this genus following the isolation of 9-isocyanopupukeanane (59) from P. varicosa.²²⁶ Evaluating the prey of the nudibranch revealed that 59 was sequestered from a sponge of the Hymeniacidon genus. The mucus secreted by P. varicosa is concentrated with respect to 59 and is known to be toxic to fish and crustaceans.²²⁶ 2-Isocyanopupukeanane was later isolated from the nudibranch and again shown to be sequestered from a Hymeniacidon sp.²²⁷epi-9-Isocyanopupukeanane (60) has been reported to co-occur with 59 in P. bourgini²²⁸ (Phyllidiella rosans⁴¹²). Thiocyano derivatives of these metabolites, 61–62, have also been isolated from P. varicosa and its sponge prey Axinyssa aculeata and were noted to be moderately antibacterial.²²⁹ Subsequent studies have found a number of related nitrogenous mono-, bi- and tricyclic sesquiterpenes from Phyllidia sp.,²³⁰P. varicosa,^231,232P. ocellata,^232–234P. coelestis,²³⁵ and P. pustulosa.^{231,232,236,237}Phyllidia pustulosa has been reclassified as Phyllidiella pustulosa⁴¹³ so unsurprisingly, several studies of the animal reported as Phyllidiella pustulosa also yielded nitrogenous sesquiterpenes,^238–242 although one additionally yielded nitrogenous diterpenes,²⁴⁰ which had also been isolated previously from a sponge.²⁴⁰Phyllidiopsis krempfi also yielded nitrogenous sesquiterpenes and a sesquiterpene peroxide.²³² Potent antifouling activity has been reported for some of these compounds against barnacle larvae^{232,237,243,244} and several also possess antimalarial activity against Plasmodium falciparum.^234,239 The lipid content of Phyllidia coelestis²⁴⁵ and the fatty acid content of P. varicosa²⁴⁶ and Phyllidiella pustulosa²⁴⁶ have also been examined.

A series of cytotoxic, antifouling sesquiterpenes (63–66) was isolated from Reticulidia fungia²⁴⁷ which possess an unique carbonimidic dichloride functionality. Two of the metabolites, reticulidins A (63) and B (64) are novel to the nudibranch and were moderately cytotoxic to two HTCLs,²⁴⁷ whilst the other two metabolites (65 and 66) had previously been isolated from the sponge Pseudaxinyssa pitys.^248,249 The structural similarities to 65, and the rarity of the carbonimidic dichloride functionality, suggests that 63 and 64 are also likely sequestered from an, as yet unidentified, Pseudaxinyssa sponge.

5.5 Polyceroidea

Although only three of the five families in the Polyceroidea have been chemically examined, the superfamily represents a remarkably diverse range of dorids that have attracted considerable attention from researchers.

5.5.1 Aegiridae. Two genera (Aegires and Notodoris) comprise the Aegiridae family but only Notodoris species have been examined to date. Nudibranchs of this genus, are specialist feeders, known to prey predominantly on sponges of the genus Leucetta,²⁵⁰ and are bright yellow, matching the colour of their sponge prey. The colouration in the sponges is caused by a range of at least 13 imidazole alkaloids^250–253 of which naamidine A (67) is a representative metabolite. Many of these compounds have also been isolated from Notodoris citrina²⁵⁰ and N. gardineri²⁵¹ after feeding on Leucetta. In addition, three metabolites only present in the nudibranch were obtained; dorimidazole A (68),²⁵¹ preclathridine A (69) and clathridine B (70).²⁵² Biosynthesis of these compounds is unlikely, due to structural similarities with other sponge metabolites.²⁵¹ Some of these compounds have been found to be antiparasitic²⁵¹ and are presumably sequestered by Notodoris for this activity.²⁵¹

5.5.2 Hexabranchidae. The family Hexabranchidae contains one genus, Hexabranchus, which has predominantly, but not exclusively, yielded macrolides. Hawaiian specimens of the bright red “Spanish dancer” nudibranch, Hexabranchus sanguineus, have yielded a number of bioactive macrocyclic lactones (macrolides). Ulapualide A (71) and B (72) were isolated from egg masses, in high concentration, and were both cytotoxic (against leukaemia) and antifungal in nature.²⁵⁴ They were also detected in the nudibranch itself, although at much lower concentrations.²⁵⁴ Three additional related macrolides, ulapualides C–E were isolated from egg masses of H. sanguineus, also obtained from Hawaii. Only ulapualide C was isolated in sufficient quantity for bioactivity testing and although it exhibited cytotoxicity against several HTCLs, it was 2-4 fold less potent than ulapualides A 71 and B 72.²⁵⁵

Other macrolides have also been isolated from H. sanguineus and its egg masses from various locations. These include kabiramide C (73)^256–258 and related compounds^{256,258–260} and halichondramide derivatives^258,260,261 such as dihydrohalichondramide (74). Ulapualides, kabiramides and halichondramide derivatives possessed a range of bioactivities, including cytotoxicity^{254,255,260,261} and antifungal properties^{254,256,259,261} and the latter two macrolide families have been shown to deter feeding in both fish (Thalassoma lunare) and crabs (Dardunus megistos).²⁵⁸ Macrolides have been found at higher concentrations in egg masses than in the adult nudibranchs,^254,258 suggesting that their main function is to protect these vulnerable egg masses, although it has been postulated that the macrolides are also secreted in the mucus of H. sanguineus thus conferring protection on the adult as well.²⁵⁸ One investigation has highlighted that the bioactivity of 71 (and hence presumably that of related compounds), is derived from potent depolymerisation of the actin cytoskeleton within cells.²⁶²

Most of these macrolides are strongly suspected of being of dietary origin as they were found in both the mantle and digestive tracts of the nudibranch.²⁵⁸ Furthermore, halichondramide²⁶¹ and derivatives have been isolated from Halichondria sponges and these sponges are known prey species of H. sanguineus. Halichondramide itself, is absent in all specimens of H. sanguineus, despite being the major metabolite of the sponge,^258,261 so it has been suggested that the nudibranch is either secondarily converting halichondramide into compounds such as 74, or selectively excluding it.²⁵⁸ Since halichondramide is very cytotoxic (LD_LO in mice when injected subcutaneously is 1.4 mg kg⁻¹),²⁶¹ it may be too potent for use by H. sanguineus.

Hurghadin (75), the red pigment responsible for the colouration of H. sanguineus, has been isolated from an Egyptian specimen²⁶³ and as it is a carotenoid, it is almost certainly sequestered. The occurrence of 75 in the mantle follows an inverse relationship with the known macrolides, indicating that it may provide an alternative chemical defence, in addition to acting as a visual warning to potential predators,²⁶³ although with no bioactivity studies conducted on 75, this is only conjecture. The apocarotenoid apoastacenal was isolated from H. sanguineus egg masses from Japan²⁶⁴ as a new natural product (but known synthetic compound).²⁶⁵

H. sanguineus is a generalist feeder on many species of sponge. An adult specimen from the South China Sea was found to contain a mixture of nitrogenous sesquiterpenoids rather than macrolides,²⁶⁶ two of which were known metabolites from Axinella, Acanthella and Dysidea sponges, and two were novel.²⁶⁶ However, as both novel compounds were found in the digestive tract of the nudibranch, this suggests that they too are of dietary origin.

A Fijian specimen of H. sanguineus analysed by very sensitive NMR spectroscopy (600 MHz spectrometer with a 1 mm high-temperature superconducting cryoprobe), yielded, in addition to a number of related macrolides, two thiazole cyclic peptides, sanguinamide A (76) and sanguinamide B (77).²⁵⁹ Both 76 and 77 were present at extremely low concentrations across the whole individual (0.0023% and 0.011% dry weight respectively) suggesting that the compounds were not major components in H. sanguineus defence and likely represented metabolites of an as yet unidentified prey species.²⁵⁹ Although no dietary source has been identified for the sanguinamides, the structures suggest that cyanobacteria or tunicates may be their original source.²⁵⁹ Both 76 (ref. 267) and 77 (ref. 68) have been synthesised; synthesis of the former revised the configuration to (76). Sanguinamide B analogues were active against Pseudomonas aeruginosa, reducing twitching motility.^268,269

The implication of these studies, is that H. sanguineus can sequester a variety of metabolites to utilise in defence and can seemingly do so from a range of prey species.^259,266

5.5.3 Polyceridae. The Polyceridae is comprised of three subfamilies, the Nembrothinae, the Polycerinae and the Triophinae. The Nembrothinae contains three genera, Nembrotha, Tambja and Roboastra, all of which utilise a series of antimicrobial and cytotoxic alkaloids, the tambjamines, which are known to be sequestered metabolites.²⁷⁰Roboastra tigris, a large predatory nudibranch, feeds upon smaller nudibranchs, preferentially Tambja abdere and T. eliora, which in turn prey on the bryozoan Sessibugula translucens. Tambjamines A–D 78–81 were found in all four organisms, indicating their production by the bryozoan then sequestration and concentration up the food chain.²⁷⁰

Tambjamines have also been noted to deter feeding in fish, and thus likely act as a defence against both generalist and specialist feeders.²⁷¹ Conversely, the Tambja nudibranchs and R. tigris have developed the ability to track their respective prey through chemoreception of these same compounds.²⁷¹ In the slime trails of T. abdere, 78–81 were present at low concentrations, as well as being released in a mucus at relatively high concentrations when T. abdere was molested.^270,271R. tigris was noted to track the slime trail of T. abdere, but would sometimes break off attack if the mucus was released,²⁷⁰ indicating that the antifeeding properties of the tambjamines still affected R. tigris if their concentration was sufficient.²⁷¹ In the event that T. eliora was attacked, it would attempt to flee rather than secrete a defence, suggesting that the lower concentration of tambjamines in T. eliora prevents release of a mucus.²⁷¹ Similarly, T. eliora preferentially selected low concentrations of tambjamines (consistent with natural concentrations in S. translucens) in a Y maze, but was deterred if concentrations were high.²⁷¹ Thus the tambjamines, in addition to being an antifeedant, in Tambja species, act as a tracking pheromone at low concentrations and an alarm pheromone at higher concentrations.²⁷¹

Tambjamines, including 78–81 and new variants, have since been reisolated from T. eliora²⁷² and from a bryozoan.²⁷³ A related yellow pigment was obtained from a bacterium²⁷⁴ and a related blue tetrapyrrole pigment was isolated independently from Nembrotha kubaryana,⁹⁸ a compound ascidian,²⁷⁵ a bryozoan²⁷⁶ and from the predator-prey pairing Nembrotha spp. and the ascidian Atapozoa sp.²⁷⁷ and was identical to that produced by a mutant bacterium,²⁷⁸ indicating that it is likely of bacterial origin in all cases. Tambjamines have also been isolated from several predator-prey pairings; Tambja ceutae and the bryozoan Bugula dentata,²⁷⁹Tambja stegosauriformis and B. dentata¹³⁷ and from Nembrotha spp. and the ascidian Atapozoa sp.^277,280 Some tambjamines, especially 81, have been investigated for pharmaceutical uses as they have been shown to be cytotoxic against a number of tumour cell lines.^272,275 Proposed mechanisms for the cytotoxicity include intercalation into DNA as well as the promotion of single-strand DNA oxidative cleavage, although a lack of selectivity for tumour cells over healthy cells has been noted as a prominent obstacle to drug development.²⁸¹

The Triophinae subfamily consists of nine genera of which only two (Triopha and Limacia) have been chemically examined. Two temperate nudibranchs from this subfamily, Triopha catalinae and Limacia clavigera also feed exclusively on bryozoans.^282,283 The structurally related triophamine (82)²⁸² and limaciamine (83)²⁸³ have been isolated from T. catalinae and L. clavigera respectively. Unlike the tambjamines, triophamine 82 has been shown to be de novo synthesised by T. catalinae^284,285 and has also been isolated from a nudibranch in the closely related Polycerinae subfamily, Polycera tricolor.¹⁵² The geometry of the double bonds has been determined by total synthesis.²⁸⁶

The structural similarities between triophamine 82 and limaciamine 83 has led to speculation that 81 is also biosynthesised,²⁸³ but this has yet to be proven. As no bioactivity studies were conducted on these metabolites, their biological roles can only be speculated upon. The carotenoid triophaxanthin was first isolated as the main pigment of Triopha carpenteri²⁸⁷ (T. catalinae⁴¹⁴) but subsequent isolations from a tunicate²⁸⁸ and a cuttlefish²⁸⁹ and its presence in the gut of T. carpenteri,²⁸⁷ indicate that it is dietary in nature.

6 Cladobranchs

6.1 Aeolidioidea

The superfamily Aeolidioidea, comprises seven families but only two (Aeolidiidae and Facelinidae) have been studied. Despite this, the superfamily has attracted much attention from researchers, largely due to the genus Phyllodesmium, the only aeolid genus known not to sequester cnidocysts or nematocysts.²⁹⁰

6.1.1 Aeolidiidae. Of the nine genera comprising the Aeolidiidae family, a single species of each of three, Aeolidia, Aeolidiella (Berghia⁴¹⁵) and Spurilla has been chemically examined. When Aeolidia papillosa ate the sea anemone Anthopleura elegantissima, the alarm pheromone from A. elegantissima, anthopleurine, persisted in the nudibranch tissues for over five days. After this time, Aeolidia papillosa could evoke alarm responses in anemones without touching them, stimulating them to withdraw the tentacles and oral disk and to leave the body regions with the highest anthopleurine concentrations open to attack.²⁹¹Aeolidiella stephanieae (Berghia stephanieae⁴¹⁵) exclusively feeds on glass anemones of the Aiptasia genus which usually contain endosymbiotic dinoflagellates that provide nutrients such as fatty acids to their host. A study where breeding pairs of Berghia stephanieae were fed with either symbiotic or aposymbiotic Aiptasia pallida, revealed differences in the fatty acid profiles of their egg masses.²⁹² An Argentinian collection of a number of individuals of Spurilla sp. revealed that the prey cnidocysts in the nudibranch cerata were highly variable in type and abundance and thus from a variety of sea anemone species.²⁹³

6.1.2 Facelinidae. Facelinidae is a large family of nudibranchs comprised of 34 genera. Representatives of just four of these genera (Cratena, Hermissenda, Phidiana and Phyllodesmium) have been studied chemically. Some prenylphenols were isolated from the skin of Cratena peregrina,²⁹⁴ two of which were found previously in a brown alga²⁹⁵ and in the digestive glands of another nudibranch, Dendrodoris grandiflora,²¹² indicating their dietary origin. Chitin was found in C. peregrina (and Flabellina affinis) as intracellular granules in skin epidermal cells and the epithelial cells of the stomach.²⁹⁶ These nudibranchs prey on Cnidaria and the chitin granules are released in response to nematocysts fired by prey tentacles and form aggregates with the nematocyst tubules, thus protecting the animals from the effects of the tentacles. In granular form, the chitin maintains skin flexibility while still conferring protection.²⁹⁶

L-6-Bromohypaphorine (84) was isolated from Hermissenda crassicornis and found to be an agonist of human α7 nicotinic acetylcholine receptor.²⁹⁷ A dietary origin of 84 is likely, as the compound has been found in the sponges Pachymatisma johnstoni²⁹⁸ and Aplysina sp.,²⁹⁹ as well as the tunicate Aplidium conicum.³⁰⁰ Whilst H. crassicornis is largely considered a generalist feeder, a recent report noted a preference for colonial tunicates in its diet, especially Aplidium solidum.³⁰¹ It would therefore seem likely that the source of 84 in the nudibranch is from either A. conicum itself or another Aplidium species.

Two indole alkaloids, phidianidines A (85) and B (86), were isolated from Phidiana militaris and were cytotoxic against a number of cell lines, including C6 and HeLa tumour cells.³⁰² They remain to date, the only known marine natural products with the 1,2,4-oxadiazole core. The rarity of this backbone, coupled with the total lack of any precursor metabolite in a known prey species, precludes the possibility of P. militaris sequestering 85 and 86. Whilst de novo synthesis might therefore be suspected, isotope studies are needed for confirmation. If isotope studies fail to show evidence of biosynthesis, then an unidentified prey species must possess the metabolites, or precursors thereof, or the source could potentially be a symbiotic organism. The biological activity of 85 and 86 coupled with their structural rarity, have led to syntheses of the natural products^303–306 and analogues^307–309 for exploration of their neuroprotective effects.^303–309

The genus Phyllodesmium is the only aeolid genus known not to sequester cnidocysts or nematocysts.²⁹⁰ Without the physical defence utilised by many aeolids, it might be expected that Phyllodesmium sp. would obtain a chemical defence in its place and indeed, this has been shown to be the case.^{290,310–312} The cytotoxic diterpene trocheliophorol (87) was isolated from Phyllodesmium longicirra (P. longicirrum⁴¹⁸) and the soft coral Sarcophyton trocheliophorum, upon which it was found.³¹⁰ The cerata of the nudibranch were concentrated with 87 and were noted to detach when molested, suggesting that P. longicirra was sequestering a defensive metabolite.³¹⁰ Similarly, Phyllodesmium guamensis (P. guamense⁴¹⁶) was also found grazing upon soft corals; Sinularia maxima, S. polydactyla and an unidentified Sinularia species.³¹¹ The diterpene 11β-acetoxypukalide (88) was sequestered from these corals by P. guamense and concentrated in its cerata. Diterpene 88 deterred feeding by the predatory pufferfish Canthigaster solandri at concentrations lower than those found in the cerata.³¹¹ Four polycyclic diterpenes were isolated from a single specimen of Phyllodesmium longicirrum from the Great Barrier Reef, Australia, of which 4-oxochatancin (89) deterred feeding by Canthigaster solandri.³¹³ Although the food source of the nudibranch was not collected, it was noted that P. longicirrum usually feeds on soft coral of the Sarcophyton genus.³¹³ This, coupled with 89 possessing a planar structure the same as that of isosarcophytin from Indian Ocean S. elegans³¹⁴ but stereochemistry the same as that of chatacin³¹⁵ from an Okinawan Sarcophyton sp., means that 89 is very likely a sequestered metabolite.

Sesquiterpenes are also sequestered by Phyllodesmium aeolids. Those obtained from P. lizardensis (P. lizardense⁴¹⁷), were also found at an elevated concentration in the cerata²⁹⁰ and their dietary source was found to be a coral of the Heteroxenia genus. Although these sesquiterpenes were inactive in antifungal, antibacterial or antialgal assays and were not tested for feeding deterrence, a related compound, (+)-6-hydroxy-α-muurolene, has been found to be lethal to brine shrimp.³¹² A mixture of eight diverse sesquiterpenes was also isolated from P. magnum, including a novel asteriscane terpene (90).³¹⁶ Of the remaining seven metabolites, four were known from corals of the Sinularia genus and three were known from the brown alga Dictyopteris divaricata,³¹⁶ suggesting sequestration and that P. magnum may graze upon corals as well as brown algae. Whilst 90 was a novel natural product, a more recent study of Sinularia capillosa highlighted compounds with similar backbones,³¹⁷ indicating that 90 may be a metabolite of an unidentified Sinularia sp., or be otherwise secondarily modified, as opposed to being biosynthesised de novo.

6.2 Arminoidea

Consisting of the two families, Arminidae (seven genera) and Doridomorphidae (one genus), this superfamily has received minimal attention from natural products researchers, with only two species of the Arminidae (Armina and Dermatobranchus) being investigated to date. A series of seven briarane diterpenes, including the chlorinated example 91, was isolated from the nudibranch Armina maculata and its octocoral prey Veretillum cynomorium^318–320 as was a cembranoid, preverecynarmin (92),³²⁰ highlighting that all were of dietary origin for the nudibranch. Similarly, four related diterpenoids, ophirin (93), calicophirin B (94), 13-deacetoxyl calicophirin B (95) and 13-deacetoxy-3-deacetyl calicophirin B (96), were isolated from the nudibranch Dermatobranchus ornatus.³²¹ Of these, 94 and 95 were subsequently found within a prey species, a gorgonian of the Muricella genus.³²¹ This indicated that these compounds may be of dietary origin, with 93 and 96 being sourced from an unidentified prey species or otherwise secondarily modified.

6.3 Doridoxoidea

Of the unassigned Cladobranchs, there has been one superfamily designated, the Doridoxoidea which in turn consists of one family (the Doridoxidae) containing one genus (Doridoxa). Under former classification systems,¹² Doridoxoidea was not strictly a cladobranch, but evidence emerged to suggest that this superfamily should be included under Cladobranchia,¹⁵ and it is now classified as such.¹³ There have been no natural product studies of Doridoxoidea species, possibly as they, like the Bathydoridoidea, are polar, deep water nudibranchs and are thus considerably more difficult to collect.¹²

6.4 Fionoidea

This superfamily contains the families Fionidae and Pseudovermidae but has barely been examined. A study of Phestilla melanobranchia (Tenellia melanobranchia⁴¹⁹) (Fionidae) which feeds on the coral Tubastrea coccinea, yielded the known sponge alkaloids, 4′-de-N-methylaplysinopsin (97) and 6-bromo-4′-de-N-methylaplysinopsin (98), which were also obtained from the prey coral.³²² A study of Phestilla sibogae (Tenellia sibogae⁴²⁰) highlighted that the nudibranch can track its coral prey through chemoreception using its rhinophores⁴ but the study did not involve isolating any metabolites that could potentially be implicated. The researchers noted that the rhinophores responded to various amino acids, with the most intense responses to aspartic and glutamic acids,⁴ so suggested that P. sibogae tracked corals through amino acids.⁴ However, as no natural product investigations have been conducted, the possibility remains that the nudibranch may detect and respond to secondary metabolites in corals that it wishes to sequester. After all, tracking of metabolites is not unprecedented in nudibranchs, as the response of Roboastra tigris to the tambjamines highlights (see Section 5.5.3 above).²⁷⁰

6.5 Flabellinoidea

The Flabellinoidea is divided into two families, the Flabellinidae (eight genera including Flabellina and Flabellinopsis) and the Notaeolidiidae (one genus) but there has been very little chemical research into these nudibranchs. As noted above, chitin was found in skin epidermal cells and the epithelial cells of the stomach of Flabellina affinis as intracellular granules.²⁹⁶F. affinis was also noted to feed and lay its eggs upon several species of hydroid of the Eudendrium genus and to contain the same polyhydroxylated sterols as the hydroid but only the major component of this sterol mix was isolated and only from the hydroid.³²³ The ubiquitous carotenoid astaxanthin³²⁴ was found in Flabellinopsis iodina and its sponge diet.²⁸⁷

6.6 Tritonioidea

Nudibranchs of this superfamily, sometimes referred to as dendronotoids, are further separated into nine families, of which only two, the Tethydidae (two genera) and the Tritoniidae (nine genera) have received any attention from natural product chemists.

6.6.1 Tethydidae. Tethydidae, comprises two genera; (Melibe and Tethys) and natural product biosynthesis has been observed in both.^325,326 Nudibranchs of this family are unusual in that they do not feed on sessile, benthic organisms, but rather on free swimming planktonic crustaceans.^7,325 They have developed an extended oral veil which acts as a net to sweep up their prey, and are noted to swim freely in the water column.³²⁵Melibe leonina is found along the entire coastline of Northern America and is noted to have a fruity aroma.³²⁷ Degraded terpenoids, 2,6-dimethyl-5-heptenoic acid (a known synthetic compound)³²⁸ and 2,6-dimethyl-5-heptenal³²⁵ (99), were isolated from the nudibranch, of which 99 was found to be the source of this scent. De novo synthesis was confirmed via stable isotope feeding studies.³²⁵ Whilst no bioactivity studies of 99 were conducted, a potential sea star predator (Pyncnopodia helianthoides) was repelled upon contact with M. leonina.³²⁵ The complete mitochondrial genome of M. leonina has recently been determined.^329,330

Prostaglandin (PG) lactones have been isolated from the related species Tethys fimbria^326,331 which is known to synthesise some of these de novo.^326,332 These are synthesised from, and act as protected forms of, the free PG acids,³²⁶ compounds known to promote a range of hormonal responses in many organisms. PG molecules are classified by the structure of their central ring system, donated by a letter, which indicates the membrane protein that the PG will interact with.³³³ A subscript indicates the number of double bonds present in the aliphatic side chains. T. fimbria synthesises and stores PG lactones of the E class, PGE₂-1,15-lactone (100) and PGE₃-1,15-lactone (101), in its cerata.³²⁶ When molested, T. fimbria emits a mucus from its body and cerata and if the cerata become detached, they can continue to release mucus for a number of hours. When molested, the nudibranch converted 100 and 101 back to the free acid forms PGE₂ (102) and PGE₃ (103) respectively,³²⁶ suggesting that the free acid forms 102 and 103 were being utilised in the nudibranch's defence. PG-lactones of the F series have also been detected in T. fimbria (not pictured, but 9(S)-hydroxy derivatives of 100 and 101)³³² but are only present in the ovotestis (hermaphroditic gland of the nudibranch) and not the cerata, indicating that they may play a role in reproduction as opposed to defence.³³²

6.6.2 Tritoniidae. The most diverse of the Tritinoidea families is Tritoniidae. Comprising nine genera, these nudibranchs are noted to sequester metabolites from their prey.^334,335 The macrolactone amphidinolide P (104), was isolated from the octocoral Stragulum bicolor and its nudibranch predator Marionia limceana from Brazil.³³⁵ Prior to this isolation, 104 has only been isolated from laboratory cultures of the marine dinoflagellate Amphidinium sp.³³⁶ but S. bicolor appears to host a diverse but so far uncultivable dinoflagellate community and 104 was present in much higher concentration in the octocoral than was reported in Amphidinium sp. Although 104 was passed up trophic levels to Marionia limceana, no evidence was presented that it was actively sequestered and despite it having been reported as possessing moderate cytotoxicity,³³⁶104 was found to be inactive when tested against the HCT-116 cell line in this study.³³⁵

A series of six terpenoid metabolites has been isolated from two nudibranch populations of Tochuina tetraquetra³³⁷ (T. gigantea⁴²³). Specimens from Port Hardy, British Columbia were noted to contain tochuinyl acetate (105), dihydrotochuinyl acetate (106), rubifolide (107) and pukalide (108). Analysis of a prey species, the soft coral Gersemia rubiformis, present at Port Hardy, indicated that 105 and 106 were minor metabolites of this species. Rubifolide 107 had been previously detected in G. rubiformis.³³⁸ Whilst a known source of 108 was not found at Port Hardy, it has been previously reported from tropical corals, Sinularia,³³⁹ and gorgonians of the genus Lophogorgia.³³⁷ A second collection of T. tetraquetra, from Bamfield, British Columbia, revealed the diterpenoid ptilosarcenone (109) and its butanoate analogue (110).³³⁷ The sea pen Ptilosarcus gurneyi, a prey species of T. tetraquetra in Bamfield waters, is known to produce 109.³⁴⁰ As it appears that the nudibranch sequesters natural products from various prey species,³³⁷ it is therefore not surprising that the geographically separated populations possess different compositions of natural products.

Tritonia hamnerorum was observed on its prey, the sea fan Gorgonia ventalina at very high densities, in the Florida Keys, U.S.A., resulting in the isolation of the furano-germacrene julieannafuran (111) from both sea fan and nudibranch. T. hamnerorum concentrated 111 relative to other sea fan metabolites and 111 either significantly reduced or entirely prevented reef fish feeding on the nudibranch in both natural and laboratory environments.³⁴¹

1-O-Hexadecyl glycerol (chimyl alcohol) was detected in the Antarctic nudibranch Tritoniella belli and its usual prey, the stoloniferan coral Clavularia frankliniana. The common predatory Antarctic sea star Odontaster validus, showed feeding deterrence to both T. belli mantle tissue and to chimyl alcohol.³⁴² A series of sesquiterpenes, tritoniopsins A–D (112–115), were isolated from the nudibranch Tritoniopsis elegans and its soft coral prey Cladiella krempfi.³³⁴ The major metabolites in both species were found to be 112 and 113 but the relative ratio of these was inverted in the predator-prey pair, such that 112 dominated the T. elegans extract and 113 dominated the extract of C. krempfi. Tritoniopsin A 112 was concentrated within the mantle tissue of T. elegans, to a greater degree than were 113–115, suggesting that 112 plays a defensive role for the nudibranch and that it was preferentially sequestered from the coral. No bioassays were conducted for 112, but weak to moderate cytotoxicity was observed for tritoniopsin B 113 against a selection of rat cell lines.³³⁴

6.7 Unassigned families

6.7.1 Charcotiidae. Only three of the nine unassigned families have been subject to natural products investigations. The first, Charcotiidae, contains three genera (Charcotia, Leminda and Pseudotritonia) and only the Antarctic nudibranch Charcotia granulosa and the tropical nudibranch Leminda millecra have been examined chemically. Granuloside (116), a homosesterterpene lactone, was isolated from the Antarctic nudibranch Charcotia granulosa.³⁴³ Whilst sesterterpenes are known in some dorid nudibranchs, granuloside (116) is the only known linear homosesterterpene in all of nature.³⁴³ It was isolated from the lipophilic extract of the external part of the nudibranch and was absent in the gut and the digestive gland. Additionally, neither 116 nor any related compound was found in the specialist prey of the nudibranch, the bryozoan Beania erecta, strongly suggesting a de novo biosynthetic origin.³⁴⁴

L. millecra is known to sequester sesquiterpenes.^345,346 Millecrones A (117) and B (118), as well as millecrols A (119) and B (120), were isolated from a South African population of L. millecra.³⁴⁵ Analysis of the nudibranch's stomach contents revealed spicules from the soft corals Alcyonium foliatum, A. valdiviae and Capnella thyrsoidea.³⁴⁵ Limited antimicrobial activity was noted for 117, 119 and 120. A separate analysis of another South African population of L. millecra and its local prey, identified the dietary origin of 117 and 118.³⁴⁶ GC analysis indicated that 117 originated from the soft coral Alcyonium fauri, whilst 118 was from the gorgonian Leptogorgia palma.³⁴⁶ At least eleven other sesquiterpenes were detected (structures not shown) in the latter study by GC. These were also expected to be of dietary origin, from various octocoral prey species, due to the backbone common with typical octocoral metabolites.³⁴⁶ Some of the triprenylated toluquinones and toluhydroquinones obtained from Leminda millecra induced apoptosis in oesophageal cancer cell lines.³⁴⁷

6.7.2 Dotidae. Members of the Dotidae family (comprising four genera) have been reported to feed exclusively on hydrozoans.³⁴⁸ Only one species from this family has been chemically studied. A Spanish collection of Doto pinnatifida yielded dotofide (121), a guanidine-interrupted terpenoid³⁴⁸ but 121 was not found in the hydrozoan Nemertesia antennina, the exclusive prey of D. pinnatifida, suggesting the likelihood of de novo biosynthesis.³⁴⁸ A new species, D. carinova has recently been described and compared with D. antarctica but as yet, no chemistry has been reported for either species.³⁴⁹

6.7.3 Proctonotidae. Of the five genera comprising the Proctonotidae family, only Janolus has been studied. The tripeptide janolusimide (122) was isolated from the Mediterranean nudibranch Janolus cristatus.³⁵⁰ Janolusimide (122) was toxic to mice (LD 5 mg kg⁻¹; i.p.) and antagonism of its neurotoxic action at lower concentrations by atropine suggested that 122 affects the acetylcholine receptors.³⁵⁰ More recently, an N-methyl analogue of 122, janolusimide B (123) was isolated from a New Zealand population of the bryozoan, Bugula flabellata³⁵¹(Bugulina flabellata³⁵²) from Stewart Island. This bryozoan is native to the British Isles but has since spread to Australasia, the western coastline of the U.S.A. and throughout the Atlantic and Mediterranean. In New Zealand at least, it is considered a fouling “weed” for all intents and purposes.³⁵³ The structural similarity between 122 and 123 suggested an ecological interaction between J. cristatus and B. flabellata.³⁵¹ Indeed, J. cristatus is known to feed predominantly on bryozoans, including B. flabellata (Fig. 3).³⁵⁴ However, as 122 was never identified in the bryozoan, and 123 was never found in the nudibranch, it was unclear if J. cristatus sequestered 122 from its diet or secondarily modified it from 123.

Fig. 3 Janolus cristatus feeding on Bugulina flabellata in the Shetland Islands. Photograph courtesy of Joanne Porter.

A subsequent investigation into a different New Zealand population of B. flabellata found evidence of both 122 and 123 within the bryozoan.³⁵⁵ Small scale extraction of New Zealand collections of Janolus novozealandicus and Bugulina flabellata and screening of the extracts by tandem liquid chromatography mass spectrometry (LCMS) indicated the presence of both 122 and 123 and suggested the presence of a series of at least six related tripeptides. The relative abundance of 122 and 123 is inverted within J. novozealandicus (122 dominant) with respect to B. flabellata (123 dominant) suggesting preferential uptake of 122. An egg mass of J. novozealandicus was also extracted and screened and of the two tripeptides, only 122 was found (as the main metabolite), highlighting its potential role as a defensive compound,³⁵⁵ although further studies are needed to confirm this.

7 Nematocysts and zooxanthellae

Sequestration of whole organisms (zooxanthellae) or nematocysts by some aeolid nudibranchs has been given its own section as, strictly speaking, it is not natural products chemistry. Nevertheless these processes deserve a brief mention. As has already been noted, virtually all aeolid species except those of the Phyllodesmium genus are known to sequester the stinging cells from their cnidarian prey.^9,290 The nematocysts are taken up by the aeolids and transported to the tips of cerata where multiple cells are incorporated into a single cnidosac.⁹ The use of a cnidosac is almost certainly in defence, as aeolids which have had their cerata removed are considerably more likely to be consumed. In most species, the cerata are also known to regenerate, indicating that they serve an important function to the aeolids.⁹ The exact mechanisms by which the nudibranch is able to protect itself from the sting of these cells are not fully understood.^9,356 The presence of epithelial linings within the viscera has been observed and noted to absorb the damage from a discharged nematocyst, thus protecting the nudibranch.⁹ Application of aeolid mucus to the sea anemone Anthopleura elegantissima lowered the number of nematocyst discharges, with respect to mucus of other nudibranchs.⁹ This highlighted that aeolids used multiple strategies to cope with the stinging cells. A relatively recent study tested the hypothesis that aeolids transfer immature nematocysts to the cnidosacs and then allow them to mature, thus avoiding the potential damage caused by transporting a charged nematocyst.³⁵⁶ Utilising a fluorescent dye, researchers were able to show that there was a change in pH as the nematocysts were incorporated into the cnidosacs. As an accumulation of protons is necessary for nematocysts to mature, this supported the hypothesis that aeolids themselves were maturing the nematocysts.³⁵⁶

Whilst sequestration of nematocysts is common in all aeolids except those of Phyllodesmium, an interesting symbiotic relationship has been observed in a number of cladobranchs. Zooxanthellae, photosynthetic dinoflagellates, are frequently found within the cells of cladobranchs' branched digestive gland.³⁵⁷ This symbiosis has been observed in a number of Phyllodesmium species and may highlight why this genus does not sequester nematocysts.^357,358 Aeolids of the Phestilla (Tenellia^419,420) (Fionidae family) and Pteraeolidia (Facelinidae family) have also been noted to possess zooxanthellae symbionts. A number of non-aeolid cladobranchs also obtain these symbiotic organisms, namely the dendronotoids of Melibe and Doto genera as well as Doridomorpha gardineri (Doridomorphidae family) and Pinufius rebus (unassigned Cladobranchia, Superfamily Arminoidea, Family Pinufidae).^357,358 The advantages offered to these cladobranchs are obvious, in that the transfer of (primary) metabolites can enhance survivability if food is scarce.³⁵⁷ Interestingly, there have been no reported cases of zooxanthellae symbiosis within any of the dorid nudibranchs. This is presumably because their unbranched digestive gland and lack of cerata (or similar projections) limits the available surface area for zooxanthellae photosynthesis.

8 Conclusions

The natural products chemistry of nudibranchs is extensive and well documented. Terpenes and sesquiterpenes in particular, dominate those metabolites that have been isolated, although virtually all classes of natural products are represented. The sequestering of these metabolites from prey species is seemingly the most common source of nudibranch natural products. Indeed, metabolites have been sequestered from sponges, corals, tunicates, gorgonians and bryozoans. Whilst de novo synthesis is rarer than sequestration of natural products, it can still be found throughout all of Nudibranchia. Secondary modification is also suspected in a number of cases. Unsurprisingly, terpenes still dominate the types of structures observed.

It is certainly true that dorids have received considerably more research attention than their cladobranch counterparts. This is perhaps due to a perceived notion that cladobranchs, especially aeolids, utilise a physical defence (cnidosacs) over a chemical one. Furthermore, many dorids are known to feed exclusively on sponges, and sponges themselves are known to be the most prolific producers of marine natural products,^36c so the natural products chemistry of those dorids that feed on sponges might also be expected to be rich in metabolites. However, as this review has highlighted, natural products are found within all branches of Cladobranchia (that have been studied). De novo synthesis is known within the dendronotoids and suspected in some aeolids. There have even been reports of unprecedented novelty, including the first examples of marine metabolites with a 1,2,4-oxadiazole core (85 and 86)³⁰² as well as the first known linear homosesterterpene (116).³⁴³ These examples challenge the historical notion that the dorids represent the greatest source of nudibranch natural products. In light of this, more investigations into cladobranchs should be conducted moving forwards.

Even within the dorid nudibranchs, there has been an uneven allocation of research attention. Certainly the Doridoidea are well represented but many of these investigations are from species of a single genus, Chromodoris. Similarly, Hexabranchus sanguineus, has received considerable attention but no reports into other Hexabranchus species have been carried out. This preference of some superfamilies and some species over others means that there are still many nudibranchs to be properly studied. Perhaps the biggest message to be gained from this review, is that very few natural products have proven ecological roles. Indeed in some cases, only structures have been reported with absolutely no consideration for their bioactivity. Of all the metabolites presented within this review for which an ecological role is known or suspected, all but the PGF-lactones isolated from Tethys fimbria, are used in defence. Although the defensive capabilities of natural products are assumed in many cases, very few have been shown to actually deter predation. However, proving a chemical ecological role is not a simple task. Typically this requires dissection of the nudibranch into its mantle, viscera and other parts, instead of whole organism extraction, as well as conducting a number of antifeeding tests and bioassays. In many cases, natural product investigations are pharmaceutically driven and the chemical ecology is only of secondary interest. Thus, extensive ecological studies are relatively rare in the literature, given the time and amount of metabolite required to adequately perform these experiments.

9 Conflicts of interest

There are no conflicts to declare.

10 Acknowledgements

We thank Tracey Bates, Rex Fairweather and Joanne Porter for the photographs provided for this review, Tracey Bates and Rex Fairweather for helpful discussions and the anonymous reviewers for their helpful suggestions which have improved this manuscript.

11 References

1. R. C. Anderson, Int. Zoo Yb., 1995, 34, 65–70,  DOI:10.1111/j.1748-1090.1995.tb00659.x.
2. S. Gofas, Nudibranchia In MolluscaBase, 2017, 15 May, 2017, accessed through: World Register of Marine Species (WoRMS) at, http://www.marinespecies.org/aphia.php?p=taxdetails%26id=1762.
3. B. E. Picton and C. C. Morrow, Nudibranchs of the British Isles, 1996, http://www.seaslug.org.uk/nudibranchs/index.html, accessed 7 January, 2016.
4. B. F. Murphy and M. G. Hadfield, Comp. Biochem. Physiol., 1997, 118A, 727–735,  DOI:10.1016/S0300-9629(97)00014-5.
5. G.-P. Hu, J. Yuan, L. Sun, Z.-G. She, J.-H. Wu, X.-J. Lan, X. Zhu, Y.-C. Lin and S.-P. Chen, Mar. Drugs, 2011, 9, 514–525,  DOI:10.3390/md9040514.
6. D. J. Faulkner and M. T. Ghiselin, Mar. Ecol.: Prog. Ser., 1983, 13, 295–301,  DOI:10.3354/meps013295.
7. G. Cimino and M. T. Ghiselin, Chemoecology, 1999, 9, 187–207,  DOI:10.1007/s000490050052.
8. M. Carbone, M. Gavagnin, M. Haber, Y.-W. Guo, A. Fontana, E. Manzo, G. Genta-Jouve, M. Tsoukatou, W. B. Rudman, G. Cimino, M. T. Ghiselin and E. Mollo, PLoS One, 2013, 8, e62075.
9. P. G. Greenwood, Toxicon, 2009, 54, 1065–1070,  DOI:10.1016/j.toxicon.2009.02.029.
10. H. Wägele and R. C. Willan, Zool. J. Linn. Soc., 2000, 130, 83–181,  DOI:10.1111/j.1096-3642.2000.tb02196.x.
11. M. Schrödl, H. Wägele and R. C. Willan, Zool. Anz., 2001, 240, 83–97,  DOI:10.1078/0044-5231-00008.
12. P. Bouchet, J. Fryda, B. Hausdorf, W. Ponder, A. Valdés and A. Warén, Malacologia, 2005, 47, 261–263.
13. WoRMS, http://www.marinespecies.org/aphia.php?p=taxdetails%26id=1762, accessed 15 May, 2017.
14. J. A. Goodheart, A. L. Bazinet, A. G. Collins and M. P. Cummings, R. Soc. Open Sci., 2015, 2, 150196,  DOI:10.1098/rsos.150196.
15. J. Mahguib and A. Valdés, Polar Biol., 2015, 38, 1369–1377,  DOI:10.1007/s00300-015-1700-5.
16. A. W. W. van Wyk and M. T. Davies-Coleman, Tetrahedron, 2007, 63, 12179–12184.
17. M. J. Garson, Chem. Rev., 1993, 93, 1699–1733,  DOI:10.1021/cr00021a003.
18. D. J. Faulkner, T. F. Molinski, R. J. Anderson, E. J. Dumdei and E. Dilip de Silva, Comp. Biochem. Physiol., Part B: Biochem. Mol. Biol., 1990, 97, 233–240.
19. P. Karuso, Bioorg. Mar. Chem., 1987, 1, 31–60,  DOI:10.1007/978-3-642-72726-9_2.
20. P. Proksch, Biol. Unserer Zeit, 1991, 21, 26–30,  DOI:10.1002/biuz.19910210110.
21. G. Cimino, A. Fontana and M. Gavagnin, Curr. Org. Chem., 1999, 3, 327–372.
22. P. Proksch, Toxicon, 1994, 32, 639–655,  DOI:10.1016/0041-0101(94)90334-4.
23. G. Cimino and G. Sodano, in Sponges Time Space, Proc. 4th Int. Porifera Congr, ed. R. W. M. Van Soest, T. M. G. Van Kempen and J.-C. Braekman, 1994, pp. 459–472.
24. M. Ishibashi, Y. Yamaguchi and Y. J. Hirano, in Biomaterials from Aquatic and Terrestrial Organisms, ed. M. Fingerman and R. Nagabhushanam, 2006, pp. 513–535.
25. G. Cimino, M. L. Ciavatta, A. Fontana and M. Gavagnin, in Bioactive Compounds from Natural Sources, ed. C. Tringali, 2001, pp. 577–637.
26. M. Gavagnin and A. Fontana, Curr. Org. Chem., 2000, 4, 1201–1248,  DOI:10.2174/1385272003375798.
27. R. J. Andersen, E. D. De Silva, E. J. Dumdei, P. T. Northcote, C. Pathirana and M. Tischler, in Recent Advances in Phytochemistry, Biochem. Mevalonic Acid Pathway Terpenoids, 1990, vol. 24, pp. 265–282.
28. M. J. Garson, J. S. Simpson, A. E. Flowers and E. J. Dumdei, Bioactive Natural Products (Part B), in Studies in Natural Products Chemistry, 2000, vol. 21, pp. 329–372.
29. M. J. Garson and J. S. Simpson, Nat. Prod. Rep., 2004, 21, 164–179,  10.1039/b302359c.
30. T. Grkovic, D. R. Appleton and B. R. Copp, Chem. N.Z., 2005, 69, 12–15.
31. M. J. Garson, Prog. Mol. Subcell. Biol., 2006, 43(Molluscs), 159–174.
32. M. J. Garson, Chem. Aust., 2004, 71, 16–18.
33. R. J. Andersen, K. Desjardine and K. Woods, Prog. Mol. Subcell. Biol., 2006, 43(Molluscs), 277–301.
34. T. Miyamoto, Prog. Mol. Subcell. Biol., 2006, 43(Molluscs), 199–214.
35. M. T. Davies-Coleman, Prog. Mol. Subcell. Biol., 2006, 43(Molluscs), 133–157.
36. (a) J. W. Blunt, B. R. Copp, R. A. Keyzers, M. H. G. Munro and M. R. Prinsep, Nat. Prod. Rep., 2017, 34, 235–294,  10.1039/C6NP00124F; (b) J. W. Blunt, B. R. Copp, R. A. Keyzers, M. H. G. Munro and M. R. Prinsep, Nat. Prod. Rep., 2016, 33, 382–431,  10.1039/C5NP00156K; (c) J. W. Blunt, B. R. Copp, R. A. Keyzers, M. H. G. Munro and M. R. Prinsep, Nat. Prod. Rep., 2015, 32, 116–211,  10.1039/C4NP00144C; (d) J. W. Blunt, B. R. Copp, R. A. Keyzers, M. H. G. Munro and M. R. Prinsep, Nat. Prod. Rep., 2014, 31, 160–258,  10.1039/c3np70117d; (e) J. W. Blunt, B. R. Copp, R. A. Keyzers, M. H. G. Munro and M. R. Prinsep, Nat. Prod. Rep., 2013, 30, 237–323,  10.1039/C2NP20112G; (f) J. W. Blunt, B. R. Copp, R. A. Keyzers, M. H. G. Munro and M. R. Prinsep, Nat. Prod. Rep., 2012, 29, 144–222,  10.1039/C2NP00090C.
37. I. W. Mudianta, A. M. White, Suciati, P. L. Katavic, R. R. Krishnaraj, A. E. Winters, E. Mollo, K. L. Cheney and M. J. Garson, Pure Appl. Chem., 2014, 86, 995–1002,  DOI:10.1515/pac-2013-1111.
38. V. T. Dang, K. Benkendorff, T. Green and P. Speck, J. Virol., 2015, 89, 8114–8118,  DOI:10.1128/JVI.00287-15.
39. N. V. Zhukova, Mar. Drugs, 2014, 12, 4578–4592,  DOI:10.3390/md12084578.
40. K. Iken, C. Avila, M. L. Ciavatta, A. Fontana and G. Cimino, Tetrahedron Lett., 1998, 39, 5635–5638,  DOI:10.1016/S0040-4039(98)01095-8.
41. C. Avila, K. Iken, A. Fontana and G. Cimino, J. Exp. Mar. Biol. Ecol., 2000, 252, 27–44,  DOI:10.1016/S0022-0981(00)00227-6.
42. MarinLit, The Royal Society of Chemistry, http://pubs.rsc.org/marinlit/, accessed 29 June 2017.
43. E. Manzo, M. Carbone, E. Mollo, C. Irace, A. Di Pascale, Y. Li, M. L. Ciavatta, G. Cimino, Y.-W. Guo and M. Gavagnin, Org. Lett., 2011, 13, 1897–1899,  DOI:10.1021/ol200377w.
44. G. Nuzzo, M. L. Ciavatta, R. Kiss, V. Mathieu, H. Leclercqz, E. Manzo, G. Villani, E. Mollo, F. Lefranc, L. D'Souza, M. Gavagnin and G. Cimino, Mar. Drugs, 2012, 10, 1799–1811,  DOI:10.3390/md10081799.
45. A. Rudi, Z. Stein, S. Green, I. Goldberg, Y. Kashman, Y. Benayahu and M. Schleyer, Tetrahedron Lett., 1994, 35, 2589–2592,  DOI:10.1016/S0040-4039(00)77179-6.
46. S. W. Ayer and R. J. Andersen, Tetrahedron Lett., 1982, 23, 1039–1042,  DOI:10.1016/S0040-4039(00)87016-1.
47. M. Gavagnin, N. Ungur, E. Mollo, J. Templado and G. Cimino, Eur. J. Org. Chem., 2002, 1500–1504,  DOI:10.1002/1099-0690(200205)2002:9<1500::AID-EJOC1500>3.0.CO;2-D.
48. J. Kubanek, D. J. Faulkner and R. J. Anderson, J. Chem. Ecol., 2000, 26, 377–389,  DOI:10.1023/A:1005405304862.
49. D. L. Burgoyne, E. J. Dumdei and R. J. Andersen, Tetrahedron, 1993, 49, 4503–4510,  DOI:10.1016/S0040-4020(01)81280-1.
50. J. E. Thompson, R. P. Walker, S. J. Wratten and D. J. Faulkner, Tetrahedron, 1982, 38, 1865–1873,  DOI:10.1016/0040-4020(82)80035-5.
51. J. Hellou, R. J. Anderson and J. E. Thompson, Tetrahedron, 1982, 38, 1875–1879,  DOI:10.1016/0040-4020(82)80036-7.
52. K. Gustafson, R. J. Andersen, C. H. He and J. Clardy, Tetrahedron Lett., 1985, 26, 2521–2524,  DOI:10.1016/S0040-4039(00)98826-9.
53. M. Tischler, R. J. Andersen, M. I. Choudhary and J. Clardy, J. Org. Chem., 1991, 56, 42–47,  DOI:10.1021/jo00001a010.
54. J. Daoust, A. Fontana, C. E. Merchant, N. J. de Voogd, B. O. Patrick, T. J. Kieffer and R. J. Andersen, Org. Lett., 2010, 12, 3208–3211,  DOI:10.1021/ol101151f.
55. J. Hellou, R. J. Andersen, S. Rafii, E. Arnold and J. Clardy, Tetrahedron Lett., 1981, 22, 4173–4176,  DOI:10.1016/S0040-4039(01)82096-7.
56. M. Tischler and R. J. Andersen, Tetrahedron Lett., 1989, 30, 5717–5720,  DOI:10.1016/S0040-4039(00)76179-X.
57. E. J. Dumdei, J. Kubanek, J. E. Coleman, J. Pika, R. J. Anderson, J. R. Steiner and J. Clardy, Can. J. Chem., 1997, 75, 773–789,  DOI:10.1139/v97-094.
58. J. Kubanek, E. I. Graziani and R. J. Andersen, J. Org. Chem., 1997, 62, 7239–7246,  DOI:10.1021/JO970695V.
59. A. Fontana, M. Gavagnin, E. Mollo, E. Trivellone, J. Ortea and G. Cimino, Comp. Biochem. Physiol., Part B: Biochem. Mol. Biol., 1995, 111, 283–290,  DOI:10.1016/0305-0491(94)00250-X.
60. A. M. White, A. S. Dewi, K. L. Cheney, A. E. Winters, J. T. Blanchfield and M. J. Garson, Nat. Prod. Commun., 2016, 11, 921–924.
61. D. E. de Silva and P. J. Scheuer, Heterocycles, 1982, 17, 167–170,  DOI:10.3987/S-1982-01-0167.
62. R. Kazlauskas, Rymantas, P. T. Murphy, R. J. Wells, K. Noack, W. E. Oberhaensli and P. Schoenholzer, Aust. J. Chem., 1979, 32, 867–880,  DOI:org/10.1071/CH9790867.
63. T. Grkovic, D. R. Appleton and B. R. Copp, Chem. N. Z., 2005, 69, 12–15.
64. R. Kazlauskas, P. T. Murphy, R. J. Quinn and R. J. Wells, Aust. J. Chem., 1976, 29, 2533–2539,  DOI:10.1071/CH9762533.
65. M. B. Ksebati and F. J. Schmitz, J. Org. Chem., 1987, 52, 3766–3773,  DOI:10.1021/jo00226a008.
66. E. Mollo, M. Gavagnin, M. Carbone, Y.-W. Guo and G. Cimino, Chemoecology, 2005, 15, 31–36,  DOI:10.1007/s00049-005-0289-5.
67. M. B. Ksebati and F. J. Schmitz, J. Nat. Prod., 1988, 51, 857–861,  DOI:10.1021/np50059a007.
68. G. Cimino, S. De Stefano, A. Guerriero and L. Minale, Tetrahedron Lett., 1975, 1425–1428,  DOI:10.1016/S0040-4039(00)72159-9.
69. R. Kazlauskas, P. T. Murphy, R. J. Wells, J. J. Daly and P. Schoenholzer, Tetrahedron Lett., 1978, 4951–4954,  DOI:10.1016/S0040-4039(01)85779-8.
70. G. M. Cameron, B. L. Stapleton, S. M. Simonsen, D. J. Brecknell and M. J. Garson, Tetrahedron, 2000, 56, 5247–5252,  DOI:10.1016/S0040-4020(00)00434-8.
71. C. Charles, J. C. Braekman, D. Daloze, B. Tursch, J. P. Declercq, G. Germain and M. Van Meerssche, Bull. Soc. Chim. Belg., 1978, 87, 481–486.
72. A. Fontana, C. Avila, E. Martinez, J. Ortea, E. Trivellone and G. Cimino, J. Chem. Ecol., 1993, 19, 339–356,  DOI:10.1007/BF00993700.
73. J. E. Hochlowski and D. J. Faulkner, Tetrahedron Lett., 1981, 22, 271–274,  DOI:10.1016/0040-4039(81)80073-1.
74. G. R. Schulte and P. J. Scheuer, Tetrahedron, 1982, 38, 1857–1863,  DOI:10.1016/0040-4020(82)80034-3.
75. J. E. Hochlowski, D. J. Faulkner, L. S. Bass and J. Clardy, J. Org. Chem., 1983, 48, 1738–1740,  DOI:10.1021/jo00158a030.
76. J. E. Hochlowski, D. J. Faulkner, G. K. Matsumoto and J. Clardy, J. Org. Chem., 1983, 48, 1141–1142,  DOI:10.1021/jo00155a055.
77. R. K. Okuda and P. J. Scheuer, Experientia, 1985, 41, 1355–1356.
78. B. Carte, M. R. Kernan, E. B. Barrabee, D. J. Faulkner, G. K. Matsumoto and J. Clardy, J. Org. Chem., 1986, 51, 3528–3532,  DOI:10.1021/jo00368a025.
79. B. Terem and P. J. Scheuer, Tetrahedron, 1986, 42, 4409–4412,  DOI:10.1016/S0040-4020(01)87279-3.
80. T. F. Molinski and D. J. Faulkner, J. Org. Chem., 1986, 51, 2601–2603,  DOI:10.1021/jo00363a040.
81. T. F. Molinski, D. J. Faulkner, C. H. He, G. D. Van Duyne and J. Clardy, J. Org. Chem., 1986, 51, 4564–4567,  DOI:10.1021/jo00374a014.
82. Y. Kakou, P. Crews and G. J. Bakus, J. Nat. Prod., 1987, 50, 482–484,  DOI:10.1021/np50051a023.
83. D. G. Corley, R. Herb, R. E. Moore, P. J. Scheuer and V. J. Paul, J. Org. Chem., 1988, 53, 3644–3646,  DOI:10.1021/jo00250a053.
84. M. R. Kernan, E. B. Barrabee and D. J. Faulkner, Comp. Biochem. Physiol., Part B: Biochem. Mol. Biol., 1988, 89B, 275–278,  DOI:10.1016/0305-0491(88)90223-4.
85. S. C. Bobzin and D. J. Faulkner, J. Org. Chem., 1989, 54, 3902–3907,  DOI:10.1021/jo00277a029.
86. E. J. Dumdei, E. D. De Silva, R. J. Andersen, M. I. Choudhary and J. Clardy, J. Am. Chem. Soc., 1989, 111, 2712–2713,  DOI:10.1021/ja00189a055.
87. G. Cimino, A. Crispino, M. Gavagnin and G. Sodano, J. Nat. Prod., 1990, 53, 102–106,  DOI:10.1021/np50067a013.
88. E. Dilip de Silva, S. A. Morris, S. Miao, E. Dumdei and R. J. Andersen, J. Nat. Prod., 1991, 54, 993–997,  DOI:10.1021/np50076a011.
89. S. A. Morris, E. Dilip de Silva and R. J. Andersen, Can. J. Chem., 1991, 69, 768–771,  DOI:10.1139/v91-114.
90. M. Gavagnin, R. R. Vardaro, C. Avila, G. Cimino and J. Ortea, J. Nat. Prod., 1992, 55, 368–371,  DOI:10.1021/np50081a014.
91. K. Sakamoto, T. Miyamoto, H. Amano, R. Higuchi, T. Komori and T. Sasaki, Tennen Yuki Kagobutsu Toronkai Koen Yoshishu, 1992, 34, 455–462.
92. T. Miyamoto, K. Sakamoto, H. Amano, R. Higuchi, T. Komori and T. Sasaki, Tetrahedron Lett., 1992, 33, 5811–5814,  DOI:10.1016/0040-4039(92)89038-E.
93. R. Puliti, M. Gavagnin, G. Cimino, C. A. Mattia and L. Mazzarella, Acta Crystallogr., Sect. C: Struct. Chem., 1992, C48, 2145–2147,  DOI:10.1107/S0108270192003196.
94. J. Pika and D. J. Faulkner, Tetrahedron, 1995, 51, 8189–8198,  DOI:10.1016/0040-4020(95)00440-J.
95. T. Miyamoto, K. Sakamoto, K. Arao, T. Komori, R. Higuchi and T. Sasaki, Tetrahedron, 1996, 52, 8187–8198,  DOI:10.1016/0040-4020(96)00388-2.
96. K. McPhail and M. T. Davies-Coleman, Tetrahedron, 1997, 53, 4655–4660,  DOI:10.1016/S0040-4020(97)00198-1.
97. T. Miyamoto, K. Sakamoto, H. Amano, Y. Arakawa, Y. Nagarekawa, T. Komori, R. Higuchi and T. Sasaki, Tetrahedron, 1999, 55, 9133–9142,  DOI:10.1016/S0040-4020(99)00477-9.
98. P. Karuso and P. J. Scheuer, Molecules, 2003, 8, 1–6,  DOI:10.3390/70100001-rev.
99. K. W. Yong, A. A. Salim and M. J. Garson, Tetrahedron, 2008, 64, 6733–6738,  DOI:10.1016/j.tet.2008.05.008.
100. M. H. Uddin, M. Otsuka, T. Muroi, A. Ono, N. Hanif, S. Matsuda, T. Higa and J. Tanaka, Chem. Pharm. Bull., 2009, 57, 885–887,  DOI:10.1248/cpb.57.885.
101. M. Agena, C. Tanaka, N. Hanif, M. Yasumoto-Hirose and J. Tanaka, Tetrahedron, 2009, 65, 1495–1499,  DOI:10.1016/j.tet.2008.11.101.
102. Suciati, L. K. Lambert and M. J. Garson, Aust. J. Chem., 2011, 64, 757–765,  DOI:10.1071/CH11036.
103. P. L. Katavic, P. Jumaryatno, J. N. A. Hooper, J. T. Blanchfield and M. J. Garson, Aust. J. Chem., 2012, 65, 531–538,  DOI:10.1071/CH12010.
104. P. L. Katavic, P. Jumaryatno, J. N. A. Hooper, J. T. Blanchfield and M. J. Garson, Aust. J. Chem., 2013, 66, 1461,  DOI:10.1071/CH12010_NC.
105. K. L. Cheney, A. White, I. W. Mudianta, A. E. Winters, M. Quezada, R. J. Capon, E. Mollo and M. J. Garson, PLoS One, 2016, 11, e0145134,  DOI:10.1371/journal.pone.0145134.
106. G. Schulte, P. J. Scheuer and O. J. McConnell, Helv. Chim. Acta, 1980, 63, 2159–2167,  DOI:10.1002/hlca.19800630805.
107. C. W. Jefford, G. Bernardinelli, J. Tanaka and T. Higa, Tetrahedron Lett., 1996, 37, 159–162,  DOI:10.1016/0040-4039(95)02113-2.
108. A. K. Ghosh and Y. Wang, J. Am. Chem. Soc., 2000, 122, 11027–11028,  DOI:10.1021/ja0027416.
109. A. Gollner and J. Mulzer, Org. Lett., 2008, 10, 4701–4704,  DOI:10.1021/ol802075v.
110. Y. Kashman, A. Groweiss, R. Lidor, D. Blasberger and S. Carmely, Tetrahedron, 1985, 41, 1905–1914,  DOI:10.1016/S0040-4020(01)96553-6.
111. M. C. A. Ramirez, D. E. Williams, J. R. Gubiani, L. L. L. Parra, M. F. C. Santos, D. D. Ferreira, J. T. Mesquita, A. G. Tempone, A. G. Ferreira, V. Padula, E. Hajdu, R. J. Andersen and R. G. S. Berlinck, J. Nat. Prod., 2017, 80, 720–725,  DOI:10.1021/acs.jnatprod.6b01160.
112. J. M. Wojnar, K. O. Dowle and P. T. Northcote, J. Nat. Prod., 2014, 77, 2288–2295,  DOI:10.1021/np500549g.
113. T. F. Molinski and D. J. Faulkner, J. Org. Chem., 1987, 52, 296–298,  DOI:10.1021/jo00378a031.
114. Y. Hirayama, P. L. Katavic, A. M. White, G. K. Pierens, L. K. Lambert, A. E. Winters, H. Kigoshi, M. Kita and M. J. Garson, Aust. J. Chem., 2016, 69, 136–144,  DOI:10.1071/CH15203.
115. E. Manzo, M. Gavagnin, M. J. Somerville, S.-C. Mao, M. L. Ciavatta, E. Mollo, P. J. Schupp, M. J. Garson, Y.-W. Guo and G. Cimino, J. Chem. Ecol., 2007, 33, 2325–2336,  DOI:10.1007/s10886-007-9387-x.
116. D. Mebs, J. Chem. Ecol., 1985, 11, 713–716,  DOI:10.1007/BF00988300.
117. M. J. Somerville, E. Mollo, G. Cimino, W. Rungprom and M. J. Garson, J. Nat. Prod., 2006, 69, 1086–1088,  DOI:10.1021/np060002i.
118. M. J. Somerville, E. Mollo, G. Cimino, W. Rungprom and M. J. Garson, J. Nat. Prod., 2007, 70, 1836,  DOI:10.1021/np070553o.
119. K. W. L. Yong, I. W. Mudianta, K. L. Cheney, E. Mollo, J. Blanchfield and M. J. Garson, J. Nat. Prod., 2015, 78, 421–430,  DOI:10.1021/np500797b.
120. A. Fontana, E. Mollo, J. Ortea, M. Gavagnin and G. Cimino, J. Nat. Prod., 2000, 63, 527–530,  DOI:10.1021/np990506z.
121. G. Cimino, S. De Rosa, S. De Stefano and G. Sodano, Comp. Biochem. Physiol., Part B: Biochem. Mol. Biol., 1982, 73B, 471–474,  DOI:10.1016/0305-0491(82)90061-X.
122. S. D. Rogers and V. J. Paul, Mar. Ecol.: Prog. Ser., 1991, 77, 221–232,  DOI:10.3354/meps077221.
123. C. Avila and V. J. Paul, Mar. Ecol.: Prog. Ser., 1997, 150, 171–180,  DOI:10.3354/meps150171.
124. A. Fontana, P. Cavaliere, N. Ungur, L. D'Souza, P. S. Parameswaram and G. Cimino, J. Nat. Prod., 1999, 62, 1367–1370,  DOI:10.1021/np9900932.
125. M. Gavagnin, E. Mollo, T. Docimo, Y.-W. Guo and G. Cimino, J. Nat. Prod., 2004, 67, 2104–2107,  DOI:10.1021/np040087s.
126. I. W. Mudianta, A. M. White and M. J. Garson, Nat. Prod. Commun., 2015, 10, 865–868.
127. A. M. White, G. K. Pierens, L. C. Forster, A. E. Winters, K. L. Cheney and M. J. Garson, J. Nat. Prod., 2016, 79, 477–483,  DOI:10.1021/acs.jnatprod.5b00866.
128. L. C. Forster, A. E. Winters, K. L. Cheney, P. Dewapriya, R. J. Capon and M. J. Garson, J. Nat. Prod., 2017, 80, 670–675,  DOI:10.1021/acs.jnatprod.6b00936.
129. J. E. Hochlowski, R. P. Walker, C. Ireland and D. J. Faulkner, J. Org. Chem., 1982, 47, 88–91,  DOI:10.1021/jo00340a018.
130. S. H. Grode and J. H. Cardellina II, J. Nat. Prod., 1984, 47, 76–83,  DOI:10.1021/np50031a009.
131. J. C. Garcia-Gomez, G. Cimino and A. Medina, Mar. Biol., 1990, 106, 245–250.
132. C. Avila, G. Cimino, A. Fontana, M. Gavagnin, J. Ortea and E. Trivellone, J. Chem. Ecol., 1991, 17, 625–636,  DOI:10.1007/BF00982131.
133. G. Cimino, A. Fontana, F. Gimenez, A. Marin, E. Mollo, E. Trivellone and E. Zubia, Experientia, 1993, 49, 582–586.
134. A. Fontana, E. Trivellone, E. Mollo, G. Cimino, C. Avila, E. Martinez and J. Ortea, J. Nat. Prod., 1994, 57, 510–513,  DOI:10.1021/np50106a011.
135. M. Haber, S. Cerfeda, M. Carbone, G. Calado, H. Gaspar, R. Neves, V. Maharajan, G. Cimino, M. Gavagnin, M. T. Ghiselin and E. Mollo, Biol. Bull., 2010, 218, 181–188,  DOI:10.1086/BBLv218n2p181.
136. J. F. Cruz, H. Gaspar and G. Calado, Chemoecology, 2012, 22, 47–53,  DOI:10.1007/s00049-011-0097-z.
137. F. R. Pereira, R. G. S. Berlinck, E. Rodrigues Filho, K. Veloso, A. G. Ferreira and V. Padula, Quim. Nova, 2012, 35, 2194–2201,  DOI:10.1590/S0100-40422012001100018.
138. G. Cimino, S. De Stefano, L. Minale and E. Trivellone, Tetrahedron Lett., 1975, 3727–3730,  DOI:10.1016/S0040-4039(00)91320-0.
139. I. W. Mudianta, V. L. Challinor, A. E. Winters, K. L. Cheney, J. J. De Voss and M. J. Garson, Beilstein J. Org. Chem., 2013, 9, 2925–2933,  DOI:10.3762/bjoc.9.329.
140. A. S. Dewi, K. L. Cheney, H. H. Urquhart, J. T. Blanchfield and M. J. Garson, Mar. Drugs, 2016, 14, 198,  DOI:10.3390/md14110198.
141. S. J. Fahey and M. J. Garson, J. Chem. Ecol., 2002, 28, 1773–1785.
142. G. Guella, I. Mancini, A. Guerriero and F. Pietra, Helv. Chim. Acta, 1985, 68, 1276–1282,  DOI:10.1002/hlca.19850680523.
143. A. Fontana, C. Muniain and G. Cimino, J. Nat. Prod., 1998, 61, 1027–1029,  DOI:10.1021/NP980073K.
144. F. A. Fuhrman, G. J. Fuhrman, Y. H. Kim, L. A. Pavelka and H. S. Mosher, Science, 1980, 207, 193–195,  DOI:10.1126/science.7350655.
145. S. J. Fahey and A. R. Carroll, J. Chem. Ecol., 2007, 33, 1226–1234,  DOI:10.1007/s10886-007-9288-z.
146. M. Gavagnin, N. Ungur, F. Castelluccio, C. Muniain and G. Cimino, J. Nat. Prod., 1999, 62, 269–274,  DOI:10.1021/NP980344R.
147. E. Zubia, M. Gavagnin, A. Crispino, E. Martinez, J. Ortea and G. Cimino, Experientia, 1993, 49, 268–271.
148. N. Ungur, M. Gavagnin, E. Mollo and G. Cimino, Tetrahedron: Asymmetry, 1999, 10, 1635–1636,  DOI:10.1016/S0957-4166(99)00160-3.
149. J. Ren, P. Zhao, X. Xiao, T. Chen and B.-B. Zeng, Synthesis, 2016, 48, 4161–4166,  DOI:10.1055/s-0035-1561493.
150. Y. H. Kim, R. J. Nachman, L. Pavelka, H. S. Mosher, F. A. Fuhrman and G. J. Fuhrman, J. Nat. Prod., 1981, 44, 206–214,  DOI:10.1021/np50014a011.
151. R. J. Quinn, R. P. Gregson, A. F. Cook and R. T. Bartlett, Tetrahedron Lett., 1980, 21, 567–568,  DOI: org/10.1016/S0040-4039(01)85558-1.
152. K. Gustafson and R. J. Andersen, Tetrahedron, 1985, 41, 1101–1108,  DOI:10.1016/S0040-4020(01)96478-6.
153. M. D. Unson, C. B. Rose, D. J. Faulkner, L. S. Brinen, J. R. Steiner and J. Clardy, J. Org. Chem., 1993, 58, 6336–6343,  DOI:10.1021/jo00075a029.
154. E. Cherbuliez and K. Bernhard, Helv. Chim. Acta, 1932, 15, 978–980,  DOI:10.1002/hlca.193201501106.
155. F. A. Fuhrman, G. J. Fuhrman, R. J. Nachman and H. S. Mosher, Science, 1981, 212, 557–558,  DOI:10.1126/science.7209552.
156. R. P. Walker and D. J. Faulkner, J. Org. Chem., 1981, 46, 1475–1478,  DOI:10.1021/jo00320a046.
157. R. P. de Jesus and D. J. Faulkner, J. Nat. Prod., 2003, 66, 671–674,  DOI:10.1021/np020542p.
158. D. E. Williams, S. W. Ayer and R. J. Anderson, Can. J. Chem., 1986, 64, 1527–1529,  DOI:10.1139/v86-250.
159. J. Kubanek and R. J. Anderson, J. Nat. Prod., 1999, 62, 777–779,  DOI:10.1021/NP9804839.
160. K. V. Reddy, R. Mohanrajd, K. N. Murthy, C. Ramesh and P. Karthick, J. Coastal Life Med., 2015, 3, 582–584,  DOI:10.12980/JCLM.3.201514J65.
161. N. K. Gulavita, P. J. Scheuer and E. D. de Silva, Bioact. Compd. Mar. Org. Indo-U. S. Symp., ed. M. F. Thompson, R. Sarojini and R. Nagabhushanam, 1991, pp. 229–233.
162. W.-F. He, Y. Li, M.-T. Feng, M. Gavagnin, E. Mollo, S.-H. Mao and Y.-W. Guo, Fitoterapia, 2014, 96, 109–114,  DOI:10.1016/j.fitote.2014.04.011.
163. R.-Y. Huang, W.-T. Chen, T. Kurtan, A. Mandi, J. Ding, J. Li, X.-W. Li and Y.-W. Guo, Future Med. Chem., 2016, 8, 17–27,  DOI:10.4155/fmc.15.169.
164. A. Fontana, P. Cavaliere, S. Wahidulla, C. G. Naik and G. Cimino, Tetrahedron, 2000, 56, 7305–7308,  DOI:10.1016/S0040-4020(00)00629-3.
165. K. Charupant, K. Suwanborirux, S. Amnuoypol, E. Saito, A. Kubo and N. Saito, Chem. Pharm. Bull., 2007, 55, 81–86,  DOI:10.1248/cpb.55.81.
166. A. Kubo, Y. Kitahara and S. Nakahara, Chem. Pharm. Bull., 1989, 37, 1384–1386,  DOI:10.1248/cpb.37.1384.
167. C. Cuevas, M. Perez, A. Francesch, C. Fernandez, J. L. Chicharro, P. Gallego, M. Zarzuelo, F. De la Calle and I. Manzanares, PCT Int. Appl., WO 2000069862 A2 20001123, 2000.
168. B. J. Petek and R. L. Jones, Molecules, 2014, 19, 12328–12335,  DOI:10.3390/molecules190812328.
169. D. Castiello, G. Cimino, S. De Rosa, S. De Stefano, G. Izzo and G. Sodano, Colloq. Int. C. N. R. S., 1979, 291, 413–416.
170. D. Castiello, G. Cimino, S. De Rosa, S. De Stefano and G. Sodano, Tetrahedron Lett., 1980, 21, 5047–5050,  DOI:10.1016/S0040-4039(00)71129-4.
171. M. L. Ciavatta, G. Nuzzo, K. Takada, V. Mathieu, R. Kiss, G. Villani and M. Gavagnin, J. Nat. Prod., 2014, 77, 1678–1684,  DOI:10.1021/np500298h.
172. P. J. Krug, K. G. Boyd and D. J. Faulkner, Tetrahedron, 1995, 51, 11063–11074,  DOI:10.1016/0040-4020(95)00679-3.
173. E. I. Graziani, R. J. Andersen, P. J. Krug and D. J. Faulkner, Tetrahedron, 1996, 52, 6869–6878,  DOI:10.1016/0040-4020(96)00327-4.
174. N. Ungur, M. Gavagnin, A. Fontana and G. Cimino, Tetrahedron: Asymmetry, 1999, 10, 1263–1273,  DOI:10.1016/S0957-4166(99)00119-6.
175. R. J. Andersen and F. W. Sum, Tetrahedron Lett., 1980, 21, 797–800,  DOI:10.1016/S0040-4039(00)71508-5.
176. K. Gustafson, R. J. Andersen, M. H. M. Chen, J. Clardy and J. E. Hochlowski, Tetrahedron Lett., 1984, 25, 11–14,  DOI:10.1016/S0040-4039(01)91135-9.
177. A. Soriente, G. Sodano, K. C. Reed and C. Todd, Nat. Prod. Lett., 1993, 3, 31–35,  DOI:10.1080/10575639308043834.
178. G. Cimino, A. Crispino, M. Gavagnin, E. Trivellone, E. Zubia, E. Martinez and J. Ortea, J. Nat. Prod., 1993, 56, 1642–1646,  DOI:10.1021/np50099a033.
179. N. Ungur, M. Gavagnin and G. Cimino, Tetrahedron Lett., 1996, 37, 3549–3552,  DOI:10.1016/0040-4039(96)00609-0.
180. R. M. Young and B. J. Baker, Planta Med., 2016, 81, S1–S381.
181. A. Cutignano, W. Zhang, C. Avila, G. Cimino and A. Fontana, Eur. J. Org. Chem., 2011, 5383–5389, S5383,  DOI:10.1002/ejoc.201100552.
182. M. T. Davies-Coleman and D. J. Faulkner, Tetrahedron, 1991, 47, 9743–9750,  DOI:10.1016/S0040-4020(01)80714-6.
183. M. Gavagnin, A. De Napoli, G. Cimino, K. Iken, C. Avila and F. J. Garcia, Tetrahedron: Asymmetry, 1999, 10, 2647–2650,  DOI:10.1016/S0957-4166(99)00273-6.
184. J. A. Maschek, E. Mevers, T. Diyabalanage, L. Chen, Y. Ren, J. B. McClintock, C. D. Amsler, J. Wu and B. J. Baker, Tetrahedron, 2012, 68, 9095–9104,  DOI:10.1016/j.tet.2012.08.045.
185. M. Gavagnin, E. Trivellone, F. Castelluccio, G. Cimino and R. Cattaneo-Vietti, Tetrahedron Lett., 1995, 36, 7319–7322,  DOI:10.1016/0040-4039(95)01476-X.
186. M. Gavagnin, M. Carbone, E. Mollo and G. Cimino, Tetrahedron, 2003, 59, 5579–5583,  DOI:10.1016/S0040-4020(03)00775-0.
187. T. Diyabalanage, K. B. Iken, J. B. McClintock, C. D. Amsler and B. J. Baker, J. Nat. Prod., 2010, 73, 416–421,  DOI:10.1021/np900617m.
188. M. Gavagnin, A. De Napoli, F. Castelluccio and G. Cimino, Tetrahedron Lett., 1999, 40, 8471–8475,  DOI:10.1016/S0040-4039(99)01777-3.
189. L. Núñez-Pons and C. Avila, Nat. Prod. Rep., 2015, 32, 1114–1130,  10.1039/C4NP00150H.
190. M. Gavagnin, M. Carbone, E. Mollo and G. Cimino, Tetrahedron Lett., 2003, 44, 1495–1498,  DOI:10.1016/S0040-4039(02)02849-6.
191. K. Iken, C. Avila, A. Fontana and M. Gavagnin, Mar. Biol., 2002, 141, 101–109,  DOI:10.1007/s00227-002-0816-7.
192. M. Gavagnin, A. Fontana, M. L. Ciavatta and G. Cimino, Ital. J. Zool., 2000, 67(Suppl.), 101–109,  DOI:10.1080/11250000009356363.
193. G. Cimino, M. Gavagnin, G. Sodano, R. Puliti, C. A. Mattia and L. Mazzarella, Tetrahedron, 1988, 44, 2301–2310,  DOI:10.1016/S0040-4020(01)81739-7.
194. M. Gavagnin, N. Ungur, F. Castelluccio and G. Cimino, Tetrahedron, 1997, 53, 1491–1504,  DOI:10.1016/S0040-4020(96)01083-6.
195. L. De Petrocellis, V. Di Marzo, B. Arca, M. Gavagnin, R. Minei and G. Cimino, Comp. Biochem. Physiol., C: Comp. Pharmacol., 1991, 100C, 603–607.
196. C. Avila, M. Ballesteros, G. Cimino, A. Crispino, M. Gavagnin and G. Sodano, Comp. Biochem. Physiol., Part B: Biochem. Mol. Biol., 1990, 97B, 363–368,  DOI:10.1016/0305-0491(90)90294-4.
197. G. Cimino, A. Crispino, S. De Stefano, M. Gavagnin and G. Sodano, Experientia, 1986, 42, 1301–1302.
198. M. Porcelli, G. Cacciapuoti, A. Oliva and V. Zappia, J. Chromatogr., 1988, 440, 151–155,  DOI:10.1016/S0021-9673(00)94519-9.
199. A. C. Granato, R. G. S. Berlinck, A. Magalhaes, A. B. Schefer, A. G. Ferreira, B. De Sanctis, J. C. De Freitas, E. Hajdu and A. E. Migotto, Quim. Nova, 2000, 23, 594–599,  DOI:10.1590/S0100-40422000000500005.
200. M. Porcelli, G. Cacciapuoti, G. Cimino, M. Gavagnin, G. Sodano and V. Zappia, Biochem. J., 1989, 263, 635–640,  DOI:10.1042/bj2630635.
201. S. W. Ayer, J. Hellou, M. Tischler and R. J. Andersen, Tetrahedron Lett., 1984, 25, 141–144,  DOI:10.1016/S0040-4039(00)99824-1.
202. S. W. Ayer, R. J. Andersen, C. H. He and J. Clardy, J. Org. Chem., 1984, 49, 2653–2654,  DOI:10.1021/jo00188a036.
203. E. I. Graziani and R. J. Andersen, J. Am. Chem. Soc., 1996, 118, 4701–4702,  DOI:10.1021/JA9539305.
204. E. I. Graziani, T. M. Allen and R. J. Andersen, Tetrahedron Lett., 1995, 36, 1763–1766,  DOI:10.1016/0040-4039(95)00118-V.
205. H. H. Strain, Biol. Bull., 1949, 97, 206–209,  DOI:10.2307/1538298.
206. J. W. McBeth, Comp. Biochem. Physiol., Part B: Biochem. Mol. Biol., 1972, 41, 69–77,  DOI:10.1016/0305-0491(72)90008-9.
207. WoRMS, http://www.marinespecies.org/aphia.php?p=taxdetails%26id=851581, accessed 23 May 2017.
208. G. Cimino, S. De Rosa, S. De Stefano, G. Sodano and G. Villani, Science, 1983, 219, 1237–1238,  DOI:10.1126/science.219.4589.1237.
209. G. Cimino, S. De Rosa, S. De Stefano and G. Sodano, Experientia, 1985, 41, 1335–1336.
210. G. Cimino, G. Sodano and A. Spinelli, J. Nat. Prod., 1988, 51, 1010–1011,  DOI:10.1021/np50059a039.
211. G. Cimino, S. De Rosa, S. De Stefano and G. Sodano, Tetrahedron Lett., 1981, 22, 1271–1272,  DOI:10.1016/S0040-4039(01)90293-X.
212. G. Cimino, S. De Rosa, S. De Stefano, R. Morrone and G. Sodano, Tetrahedron, 1985, 41, 1093–1100,  DOI:10.1016/S0040-4020(01)96477-4.
213. A. Fontana, M. L. Ciavatta, T. Miyamoto, A. Spinella and G. Cimino, Tetrahedron, 1999, 55, 5937–5946,  DOI:10.1016/S0040-4020(99)00256-2.
214. Y. Sakio, Y. J. Hirano, M. Hayashi, K. Komiyama and M. Ishibashi, J. Nat. Prod., 2001, 64, 726–731,  DOI:10.1021/np000639g.
215. R. K. Okuda, P. J. Scheuer, J. E. Hochlowski, R. P. Walker and D. J. Faulkner, J. Org. Chem., 1983, 48, 1866–1869,  DOI:10.1021/jo00159a014.
216. M. Gavagnin, E. Mollo, G. Calado, S. Fahey, M. Ghiselin, J. Ortea and G. Cimino, Chemoecology, 2001, 11, 131–136,  DOI:10.1007/PL00001843.
217. C. Avila, G. Cimino, A. Crispino and A. Spinella, Experientia, 1991, 47, 306–310,  DOI:10.1007/BF01958169.
218. A. Fontana, G. Villani and G. Cimino, Tetrahedron Lett., 2000, 41, 2429–2433,  DOI:10.1016/S0040-4039(00)00135-0.
219. N. V. Zhukova and M. G. Eliseikina, Mar. Biol., 2012, 159, 1783–1794,  DOI:10.1007/s00227-012-1969-7.
220. Y. Asakawa and T. Aratani, Bull. Soc. Chim. Fr., 1976, 9–10, 1469–1470.
221. H. Gaspar, M. Gavagnin, G. Calado, F. Castelluccio, E. Mollo and G. Cimino, Tetrahedron, 2005, 61, 11032–11037,  DOI:10.1016/j.tet.2005.08.096.
222. A. Fontana, A. Tramice, A. Cutignano, G. d'Ippolito, M. Gavagnin and G. Cimino, J. Org. Chem., 2003, 68, 2405–2409,  DOI:10.1021/jo026131v.
223. M. Gavagnin, E. Mollo, F. Castelluccio, M. T. Ghiselin, G. Calado and G. Cimino, Tetrahedron, 2001, 57, 8913–8916,  DOI:10.1016/S0040-4020(01)00876-6.
224. A. Spinella, L. A. Alvarez, C. Avila and G. Cimino, Tetrahedron Lett., 1994, 35, 8665–8668,  DOI:10.1016/S0040-4039(00)78466-8.
225. H. Gaspar, A. Cutignano, T. Ferreira, G. Calado, G. Cimino and A. Fontana, J. Nat. Prod., 2008, 71, 2053–2056,  DOI:10.1021/np8004387.
226. B. J. Burreson, P. J. Scheuer, J. Finer and J. Clardy, J. Am. Chem. Soc., 1975, 97, 4763–4764,  DOI:10.1021/ja00849a053.
227. M. R. Hagadone, B. J. Burreson, P. J. Scheuer, J. S. Finer and J. Clardy, Helv. Chim. Acta, 1979, 62, 2484–2894,  DOI:10.1002/hlca.19790620742.
228. N. Fusetani, H. J. Wolstenholme and S. Matsunaga, Tetrahedron Lett., 1990, 31, 5623–5624,  DOI:10.1016/S0040-4039(00)97915-2.
229. R. A. Edrada, V. Wray and P. Proksch, J. Nat. Prod., 2003, 66, 1512–1514,  DOI:10.1021/np030237j.
230. N. K. Gulavita, E. D. De Silva, M. R. Hagadone, P. Karuso, P. J. Scheuer, G. D. Van Duyne and J. Clardy, J. Org. Chem., 1986, 51, 5136–5139,  DOI:10.1021/jo00376a015.
231. K. E. Kassühlke, B. C. M. Potts and D. J. Faulkner, J. Org. Chem., 1991, 56, 3747–3750,  DOI:10.1021/jo00011a065.
232. T. Okino, E. Yoshimura, H. Hirota and N. Fusetani, Tetrahedron, 1996, 52, 9447–9454,  DOI:10.1016/0040-4020(96)00481-4.
233. N. Fusetani, H. J. Wolstenholme, K. Shinoda, N. Asai and S. Matsunaga, Tetrahedron Lett., 1992, 33, 6823–6826.
234. A. M. White, G. K. Pierens, T. Skinner-Adams, K. T. Andrews, P. V. Bernhardt, E. H. Krenske, E. Mollo and M. J. Garson, J. Nat. Prod., 2015, 78, 1422–1427,  DOI:10.1021/acs.jnatprod.5b00354.
235. S. Jaisamut, S. Prabpai, C. Tancharoen, S. Yuenyongsawad, S. Hannongbua, P. Kongsaeree and A. Plubrukarn, J. Nat. Prod., 2013, 76, 2158–2161,  DOI:10.1021/np4007074.
236. N. Fusetani, H. J. Wolstenholme and S. Matsunaga, Tetrahedron Lett., 1991, 32, 7291–7294,  DOI:10.1016/0040-4039(91)80501-V.
237. H. Hirota, T. Okino, E. Yoshimura and N. Fusetani, Tetrahedron, 1998, 54, 13971–13980,  DOI:10.1016/S0040–4020(98)00867-9.
238. E. J. Dumdei, A. E. Flowers, M. J. Garson and C. J. Moore, Comp. Biochem. Physiol., Part A: Mol. Integr. Physiol., 1997, 118A, 1385–1392,  DOI:10.1016/S0300-9629(97)00051-0.
239. A. D. Wright, Comp. Biochem. Physiol., Part A: Mol. Integr. Physiol., 2003, 134A, 307–313,  DOI:10.1016/S1095-6433(02)00265-9.
240. E. Manzo, M. L. Ciavatta, M. Gavagnin, E. Mollo, Y.-W. Guo and G. Cimino, J. Nat. Prod., 2004, 67, 1701–1704,  DOI:10.1021/np0400961.
241. E. G. Lyakhova, S. A. Kolesnikova, A. I. Kalinovskii and V. A. Stonik, Chem. Nat. Compd., 2010, 46, 534–538,  DOI:10.1007/s10600-010-9670-x.
242. T. Jomoria, T. Shibutani, P. Ahmadi, T. Suzuka and J. Tanaka, Nat. Prod. Commun., 2015, 10, 1913–1914.
243. N. Fusetani, H. Hiroto, T. Okino, Y. Tomono and E. Yoshimura, J. Nat. Toxins, 1996, 5, 249–259.
244. K. Nishikawa, H. Nakahara, Y. Shirokura, Y. Nogata, E. Yoshimura, T. Umezawa, T. Okino and F. Matsuda, J. Org. Chem., 2011, 76, 6558–6573,  DOI:10.1021/jo2008109.
245. N. V. Zhukova, Lipids, 2007, 42, 1169–1175,  DOI:10.1007/s11745-007-3123-8.
246. A. Alqudahi, S. B. Saad, D. Susanti and N. F. Hadry, Malays. Appl. Biol., 2016, 45, 23–28.
247. J. Tanaka and T. Higa, J. Nat. Prod., 1999, 62, 1339–1340,  DOI:10.1021/np990186j.
248. S. J. Wratten and D. J. Faulkner, Tetrahedron Lett., 1978, 16, 1395–1398,  DOI:10.1016/S0040-4039(01)94554-X.
249. S. J. Wratten, D. J. Faulkner, D. Van Engen and J. Clardy, Tetrahedron Lett., 1978, 16, 1391–1394.
250. S. Carmely, M. Ilan and Y. Kashman, Tetrahedron, 1989, 45, 2193–2200.
251. K. A. Alvi, P. Crews and D. G. Loughhead, J. Nat. Prod., 1991, 54, 1509–1515,  DOI:10.1021/np50078a004.
252. K. A. Alvi, B. M. Peters, L. M. Hunter and P. Crews, Tetrahedron, 1993, 49, 329–336,  DOI:10.1016/S0040-4020(01)80302-1.
253. A. R. Carroll, B. F. Bowden and J. C. Coll, Aust. J. Chem., 1993, 46, 1229–1234,  DOI:10.1071/CH9931229.
254. J. A. Roesener and P. J. Scheuer, J. Am. Chem. Soc., 1986, 108, 846–847,  DOI:10.1021/ja00264a052.
255. S. M. Parrish, W. Yoshida, B. Yang and P. G. Williams, J. Nat. Prod., 2017, 80, 726–730,  DOI:10.1021/acs.jnatprod.6b00896.
256. S. Matsunaga, N. Fusetani, K. Hashimoto, K. Koseki and M. Noma, Tennen Yuki Kagobutsu Toronkai Koen Yoshishu, 1985, 27, 375–382.
257. S. Matsunaga, N. Fusetani, K. Hashimoto, K. Koseki and M. Noma, J. Am. Chem. Soc., 1986, 108, 847–849,  DOI:10.1021/ja00264a053.
258. J. R. Pawlik, M. R. Kernan, T. F. Molinski, M. K. Harper and D. J. Faulkner, J. Exp. Mar. Biol. Ecol., 1988, 119, 99–109,  DOI:10.1016/0022-0981(88)90225-0.
259. D. S. Dalisay, E. W. Rogers, A. S. Edison and T. F. Molinski, J. Nat. Prod., 2009, 72, 732–738,  DOI:10.1021/np8007649.
260. S. Matsunaga, N. Fusetani, K. Hashimoto, K. Koseki, M. Noma, H. Noguchi and U. Sankawa, J. Org. Chem., 1989, 54, 1360–1363,  DOI:10.1021/jo00267a024.
261. M. R. Kernan, T. F. Molinski and D. J. Faulkner, J. Org. Chem., 1988, 53, 5014–5020,  DOI:10.1021/jo00256a021.
262. E. Vincent, J. Saxton, C. Baker-Glenn, I. Moal, J. D. Hirst, G. Pattenden and P. E. Shaw, Cell. Mol. Life Sci., 2007, 64, 487–497,  DOI:10.1007/s00018-007-6427-1.
263. Y. Guo, M. Gavagnin, E. Mollo, E. Trivellone and G. Cimino, Tetrahedron Lett., 1998, 39, 2635–2638,  DOI:10.1016/S0040-4039(98)00225-1.
264. Y. Tanaka, S. Yamada and M. Sameshima, Nippon Suisan Gakkaishi, 1992, 58, 1549,  DOI:10.2331/suisan.58.1549.
265. R. E. Coman and B. C. L. Weedon, J. Chem. Soc., Perkin Trans. 1, 1975, 2529–2532.
266. W. Zhang, M. Gavagnin, Y.-W. Guo, E. Mollo, M. T. Ghiselin and G. Cimino, Tetrahedron, 2007, 63, 4725–4729,  DOI:10.1016/j.tet.2007.03.082.
267. D. S. Nielsen, H. N. Hoang, R.-J. Lohman, F. Diness and D. P. Fairlie, Org. Lett., 2012, 14, 5720–5723,  DOI:10.1021/ol3027347.
268. E. K. Singh, D. M. Ramsey and S. R. McAlpine, Org. Lett., 2012, 14, 1198–1201,  DOI:10.1021/ol203290n.
269. H. Wahyudi, W. Tantisantisom, X. Liu, D. M. Ramsey, E. K. Singh and S. R. McAlpine, J. Org. Chem., 2012, 77, 10596–10616,  DOI:10.1021/jo3017499.
270. B. Carté and D. J. Faulkner, J. Org. Chem., 1983, 48, 2314–2318.
271. B. Carté and D. J. Faulkner, J. Chem. Ecol., 1986, 12, 795–804.
272. A. C. Granato, J. H. H. L. de Oliveira, M. H. R. Seleghim, R. G. S. Berlinck, M. L. Macedo, A. G. Ferreira, R. M. da Rocha, E. Hajdu, S. Peixinho, C. O. Pessoa, M. O. Moraes and B. C. Cavalcanti, Quim. Nova, 2005, 28, 192–198,  DOI:10.1590/S0100-40422005000200005.
273. A. J. Blackman and C. Li, Aust. J. Chem., 1994, 47, 1625–1629.
274. A. Franks, P. Haywood, C. Holmstrom, S. Egan, S. Kjelleberg and N. Kumar, Molecules, 2005, 10, 1286–1291.
275. R. Kazlauskas, J. F. Marwood, P. T. Murphy and R. J. Wells, Aust. J. Chem., 1982, 35, 215–217,  DOI:10.1071/CH9820215.
276. S. Matsunaga, N. Fusetani and K. Hashimoto, Experientia, 1986, 42, 84.
277. V. J. Paul, N. Lindquist and W. Fenical, Mar. Ecol.: Prog. Ser., 1990, 59, 109–118.
278. H. H. Wasserman, D. J. Friedland and D. A. Morrison, Tetrahedron Lett., 1968, 641–644,  DOI:10.1016/S0040-4039(00)75602-4.
279. M. Carbone, C. Irace, F. Costagilola, F. Castelluccio, G. Villani, G. Calado, V. Padula, G. Cimino, J. L. Cervera, R. Santamaria and M. Gavagnin, Bioorg. Med. Chem. Lett., 2010, 20, 2668–2670,  DOI:10.1016/j.bmcl.2010.02.020.
280. N. Lindquist and W. Fenical, Experientia, 1991, 47, 504–506.
281. B. C. Cavalcnti, H. V. N. Júnior, M. H. R. Seleghim, R. G. S. Berlinck, G. M. A. Cunha, M. O. Moraes and C. Pessoa, Chem.-Biol. Interact., 2008, 174, 155–162,  DOI:10.1016/j.cbi.2008.05.029.
282. K. Gustafson and R. J. Anderson, J. Org. Chem., 1982, 47, 2167–2169,  DOI:10.1021/jo00132a036.
283. E. I. Graziani and R. J. Anderson, J. Nat. Prod., 1998, 61, 285–286,  DOI:10.1021/NP970397T.
284. E. I. Graziani and R. J. Andersen, Chem. Commun., 1996, 2377–2378,  10.1039/cc9960002377.
285. J. Kubanek and R. J. Andersen, Tetrahedron Lett., 1997, 38, 6327–6330,  DOI:10.1016/S0040-4039(97)01455-X.
286. E. Piers, J. M. Chong, K. Gustafson and R. J. Andersen, Can. J. Chem., 1984, 62, 1–5,  DOI:10.1139/v84-001.
287. J. W. McBeth, Comp. Biochem. Physiol., Part B: Biochem. Mol. Biol., 1972, 41, 55–68,  DOI:10.1016/0305-0491(72)90007-7.
288. T. Matsuno, M. Ookubo and T. Komori, J. Nat. Prod., 1985, 48, 606–613,  DOI:10.1021/np50040a015.
289. B. Czeczuga, Hydrobiologia, 1976, 51, 71–75,  DOI:10.1007/BF00007987.
290. S. Affeld, S. Kehraus, H. Wägele and G. M. König, J. Nat. Prod., 2009, 72, 298–300,  DOI:10.1021/np800583e.
291. N. R. Howe and L. G. Harris, J. Chem. Ecol., 1978, 4, 551–561,  DOI:10.1007/BF00988919.
292. M. C. Leal, C. Nunes, D. Alexandre, T. L. Silva, A. Reis, M. T. Dinis and R. Calado, Mar. Biol., 2012, 159, 1745–1751,  DOI:10.1007/s00227-012-1962-1.
293. A. Garese, S. Garcia-Matucheski, F. H. Acuna and C. Muniain, Zool. Stud., 2012, 51, 905–912.
294. M. L. Ciavatta, E. Trivellone, G. Villani and G. Cimino, Gazz. Chim. Ital., 1996, 126, 707–710.
295. S. J. Rochfort and R. J. Capon, J. Nat. Prod., 1994, 57, 849–851,  DOI:10.1021/np50108a029.
296. R. Martin, S. Hild, P. Walther, K. Ploss, W. Boland and K.-H. Tomaschko, Biol. Bull., 2007, 213, 307–315,  DOI:10.2307/25066648.
297. I. E. Kasheverov, I. V. Shelukhina, D. S. Kudryavtsev, T. N. Makarieva, E. N. Spirova, A. G. Guzii, V. A. Stonik and V. I. Tsetlin, Mar. Drugs, 2015, 13, 1255–1266,  DOI:10.3390/md13031255.
298. W. D. Raverty, R. H. Thomson and T. J. King, J. Chem. Soc., Perkin Trans. 1, 1977, 10, 1204–1211.
299. K. Kondo and J. Nishi, J. Nat. Prod., 1994, 57, 1008–1011,  DOI:10.1021/np50109a023.
300. A. Aiello, F. Borrelli and R. Capasso, Bioorg. Med. Chem. Lett., 2003, 13, 4481–4483,  DOI:10.1016/j.bmcl.2003.08.081.
301. C. Megina, T. Gosliner and J. L. Cervera, Cah. Biol. Mar., 2007, 48, 1–7.
302. M. Carbone, Y. Li, C. Irace, E. Mollo, F. Castelluccio, A. Di Pascale, G. Cimino, R. Santamaria, Y.-W. Guo and M. Gavagnin, Org. Lett., 2011, 13, 2516–2519,  DOI:10.1021/ol200234r.
303. J. T. Brogan, S. L. Stoops and C. W. Lindsley, ACS Chem. Neurosci., 2012, 3, 658–664,  DOI:10.1021/cn300064r.
304. H.-Y. Lin and B. B. Snider, J. Org. Chem., 2012, 77, 4832–4836,  DOI:10.1021/jo300449n.
305. E. Manzo, D. Pagano, M. Carbone, M. L. Ciavatta and M. Gavagnin, ARKIVOC, 2012, 220–228,  DOI:10.3998/ark.5550190.0013.919.
306. J. C. Buchanan, B. P. Petersen and S. Chamberland, Tetrahedron Lett., 2013, 54, 6002–6004,  DOI:10.1016/j.tetlet.2013.08.063.
307. C. V. Maftei, E. Fodor, P. G. Jones, M. H. Franz, G. Kelter, H. Fiebig and I. Neda, Beilstein J. Org. Chem., 2013, 9, 2202–2215,  DOI:10.3762/bjoc.9.259.
308. C.-S. Jiang, Y. Fu, L. Zhang, J.-X. Gong, Z.-Z. Wang, W. Xiao, H.-Y. Zhang and Y.-W. Guo, Bioorg. Med. Chem. Lett., 2015, 25, 216–220,  DOI:10.1016/j.bmcl.2014.11.068.
309. L. Zhang, C.-S. Jiang, L.-X. Gao, J.-X. Gong, Z.-H. Wang, J.-Y. Li, J. Li, X.-W. Li and Y.-W. Guo, Bioorg. Med. Chem. Lett., 2016, 26, 778–781,  DOI:10.1016/j.bmcl.2015.12.097.
310. J. C. Coll, B. F. Bowden, D. M. Tapiolas, R. H. Willis, P. Djura, M. Streamer and L. Trott, Tetrahedron, 1985, 41, 1085–1092,  DOI:10.1016/S0040-4020(01)96476-2.
311. M. Slattery, C. Avila, J. Starmer and V. J. Paul, J. Exp. Mar. Biol. Ecol., 1998, 226, 33–49,  DOI:10.1016/S0022–0981(97)00240-2.
312. R. A. Edrada, V. Wray, L. Witte, L. van Ofwegen and P. Proksch, Z. Naturforsch., C: J. Biosci., 2000, 55, 82–86,  DOI:10.1515/znc-2000-1-216.
313. A. Bogdanov, C. Hertzer, S. Kehraus, S. Nietzer, S. Rohde, P. J. Schupp, H. Waegele and G. M. König, J. Nat. Prod., 2016, 79, 611–615,  DOI:10.1021/acs.jnatprod.5b00860.
314. A. S. R. Anjaneyulu, P. M. Gowri, M. J. R. V. Venugopal, P. Sarada, M. V. R. K. Murthy, G. V. Rao, P. S. N. Murthy, C. V. Rao and G. Kumar, J. Indian Chem. Soc., 1999, 76, 651–659.
315. M. Sugano, T. Shindo, A. Sato, Y. Iijima, T. Oshima, H. Kuwano and T. Hata, J. Org. Chem., 1990, 55, 5803–5804,  DOI:10.1021/jo00310a001.
316. S.-C. Mao, M. Gavagnin, E. Mollo and Y.-W. Guo, Biochem. Syst. Ecol., 2011, 39, 408–411,  DOI:10.1016/j.bse.2011.05.018.
317. D. Chen, W. Cheng, D. Liu, L. van Ofwegen, P. Proksch and W. Lin, Tetrahedron Lett., 2014, 55, 3077–3082,  DOI:10.1016/j.tetlet.2014.03.132.
318. A. Guerriero, M. D'Ambrosio and F. Pietra, Helv. Chim. Acta, 1987, 70, 984–991,  DOI:10.1002/hlca.19870700408.
319. A. Guerriero, M. D'Ambrosio and F. Pietra, Helv. Chim. Acta, 1988, 71, 472–485,  DOI:10.1002/hlca.19880710221.
320. A. Guerriero, M. D'Ambrosio and F. Pietra, Helv. Chim. Acta, 1990, 73, 277–283,  DOI:10.1002/hlca.19900730206.
321. W. Zhang, M. Gavagnin, Y.-W. Guo, E. Mollo and G. Cimino, Chin. J. Org. Chem., 2006, 26, 1667–1672.
322. R. K. Okuda, D. Klein, R. B. Kinnel, M. Li and P. J. Scheuer, Pure Appl. Chem., 1982, 54, 1907–1914,  DOI:10.1351/pac198254101907.
323. G. Cimino, S. De Rosa, S. De Stefano and G. Sodano, Tetrahedron Lett., 1980, 21, 3303–3304,  DOI:10.1016/S0040-4039(00)78673-4.
324. R. Kuhn, J. Stene and N. A. Sorensen, Ber. Dtsch. Chem. Ges. B, 1939, 72B, 1688–1701.
325. T. Barsby, R. G. Linington and R. J. Anderson, Chemoecology, 2002, 12, 199–202,  DOI:10.1007/PL00012669.
326. G. Cimino, A. Crispino, V. Di Marzo, G. Sodano, A. Spinella and G. Villani, Experientia, 1991, 47, 56–60.
327. S. W. Ayer and R. J. Andersen, Experientia, 1983, 39, 255–256,  DOI:10.1007/BF01955289.
328. D. Barnard and L. Bateman, J. Chem. Soc., 1950, 932–936,  10.1039/JR9500000932.
329. J. L. Sevigny, L. E. Kirouac, W. K. Thomas, J. S. Ramsdell, K. E. Lawlor, O. Sharifi, S. Grewal, C. Baysdorfer, K. Curr, A. A. Naimie, K. Okamoto, J. A. Murray and J. M. Newcomb, PLoS One, 2015, 10, e0127519,  DOI:10.1371/journal.pone.0127519.
330. J. L. Sevigny, L. E. Kirouac, W. K. Thomas, J. S. Ramsdell, K. E. Lawlor, O. Sharifi, S. Grewal, C. Baysdorfer, K. Curr, A. A. Naimie, K. Okamoto, J. A. Murray and J. M. Newcomb, PLoS One, 2015, 10, e0132861,  DOI:10.1371/journal.pone.0132861.
331. G. Cimino, A. Spinella and G. Sodano, Tetrahedron Lett., 1989, 30, 3589–3592,  DOI:10.1016/S0040-4039(00)99449-8.
332. G. Cimino, A. Crispino, V. Di Marzo, A. Spinella and G. Sodano, J. Org. Chem., 1991, 56, 2907–2911,  DOI:10.1021/jo00008a056.
333. K. Tsuboi, Y. Sugimoto and A. Ichikawa, Prostaglandins Other Lipid Mediators, 2002, 68–69, 535–556,  DOI:10.1016/S0090-6980(02)00054-0.
334. M. L. Ciavatta, E. Manzo, E. Mollo, C. A. Mattia, C. Tedesco, C. Irace, Y.-W. Guo, X.-B. Li, G. Cimino and M. Gavagnin, J. Nat. Prod., 2011, 74, 1902–1907,  DOI:10.1021/np200342k.
335. T. S. Sousa, G. Nuzzo, M. C. M. Torres, N. P. Lopes, A. Cutignano, P. C. Jimenez, E. A. Santos, B. A. Gomes, A. Sardo, O. D. L. Pessoa, L. V. Costa-Lotufo and A. Fontana, Rev. Bras. Farmacogn., 2015, 25, 600–604,  DOI:10.1016/j.bjp.2015.08.010.
336. M. Ishibashi, M. Takahashi and J. Kobayashi, J. Org. Chem., 1995, 60, 6062–6066,  DOI:10.1021/jo00124a015.
337. D. E. Williams and R. J. Anderson, Can. J. Chem., 1987, 65, 2244–2247,  DOI:10.1139/v87-374.
338. D. E. Williams, R. J. Anderson, G. Van Duyne and J. Clardy, J. Org. Chem., 1987, 52, 332–335,  DOI:10.1021/jo00379a002.
339. M. G. Missakian, B. J. Burreson and P. J. Scheuer, Tetrahedron, 1975, 31, 2513–2515,  DOI:10.1016/0040-4020(75)80262-6.
340. S. J. Wratten, W. Fenical, D. J. Faulkner and J. C. Wekell, Tetrahedron Lett., 1977, 18, 1559–1562,  DOI:10.1016/S0040-4039(01)93102-8.
341. G. Cronin, M. E. Hay, W. Fenical and N. Lindquist, Mar. Ecol.: Prog. Ser., 1995, 119, 177–189,  DOI:10.3354/meps119177.
342. J. B. McClintock, B. J. Baker, M. Slattery, J. N. Heine, P. J. Bryan, W. Yoshida, M. T. Davies- Coleman and D. J. Faulkner, J. Chem. Ecol., 1994, 20, 3361–3372,  DOI:10.1007/BF02033732.
343. A. Cutignano, J. Moles, C. Avila and A. Fontana, J. Nat. Prod., 2015, 78, 1761–1764,  DOI:10.1021/acs.jnatprod.5b00378.
344. J. Moles, H. Wagele, A. Cutignano, A. Fontana and C. Avila, Mar. Biol., 2016, 163, 1–11,  DOI:10.1007/s00227-016-2831-0.
345. J. Pika and D. J. Faulkner, Tetrahedron, 1994, 50, 3065–3070,  DOI:10.1016/S0040-4020(01)81106-6.
346. K. L. McPhail, M. T. Davies-Coleman and J. Starmer, J. Nat. Prod., 2001, 64, 1183–1190,  DOI:10.1021/np010085x.
347. C. E. Whibley, K. L. McPhail, R. A. Keyzers, M. F. Maritz, V. D. Leaner, M. J. Birrer, M. T. Davies-Coleman and D. T. Hendricks, Mol. Cancer Ther., 2007, 6, 2535–2543,  DOI:10.1158/1535-7163.MCT-06-0760.
348. A. Putz, S. Kehraus, G. Diaz-Agras, H. Waegele and G. M. Koenig, Eur. J. Org. Chem., 2011, 3733–3737, S3733,  DOI:10.1002/ejoc.201100347.
349. J. Moles, H. Waegele, M. Ballesteros, A. Pujals, G. Uhl and C. Avila, PLoS One, 2016, 11, e0157941,  DOI:10.1371/journal.pone.0157941.
350. G. Sodano and A. Spinella, Tetrahedron Lett., 1986, 27, 2505–2508,  DOI:10.1016/S0040-4039(00)84569-4.
351. J. Wang, M. R. Prinsep, D. P. Gordon, M. J. Page and B. R. Copp, J. Nat. Prod., 2015, 78, 530–533,  DOI:10.1021/np500752y.
352. WoRMS, http://www.marinespecies.org/aphia.php?p=taxdetails%26id=834002, accessed 12 June 2017.
353. G. Inglis, N. Gust, I. Fitridge, O. Floerl, C. Woods, B. Hayden and G. Fenwick, Port of Tauranga: Baseline survey for non-indigenous marine species, Ministry of Agriculture and Forestry Biosecurity, New Zealand: Wellington, NZ, 2005, p. 23.
354. OPK, http://opistobranquis.info/en/guia/nudibranchia/dexiarchia/unassigned-cladobranchia/janolus-cristatus/#gsc.tab=0, accessed 29 July 2017.
355. L. Dean, M. Prinsep and R. Fairweather, Chemical Theft: The Sequestration of Natural Products by New Zealand Nudibranchs from their Bryozoan Prey, in NZIC-16, The New Zealand Institute of Chemistry Conference, Queenstown, New Zealand, August, 2016, pp. 21–24, http://www.nzic16.org/.
356. D. Obermann, U. Bickmeyer and H. Wägele, Toxicon, 2012, 60, 1108–1116,  DOI:10.1016/j.toxicon.2012.08.003.
357. H. Wägele and G. Johnsen, Org. Divers. Evol., 2001, 1, 193–210,  DOI:10.1078/1439-6092-00016.
358. I. Burghardt, K. Stemmer and H. Wägele, Org. Divers. Evol., 2008, 8, 66–76,  DOI:10.1016/j.ode.2007.01.001.
359. WoRMS, http://www.marinespecies.org/aphia.php?p=taxdetails%26id=581809, accessed 19 May, 2017.
360. WoRMS, http://www.marinespecies.org/aphia.php?p=taxdetails%26id=558499, accessed 11^th May 2017.
361. WoRMS, http://www.marinespecies.org/aphia.php?p=taxdetails%26id=558620, accessed 5 October 2017.
362. WoRMS, http://www.marinespecies.org/aphia.php?p=taxdetails%26id=565620, accessed 15 May 2017.
363. WoRMS, http://www.marinespecies.org/aphia.php?p=taxdetails%26id=597348, accessed 11^th May 2017.
364. WoRMS, http://www.marinespecies.org/aphia.php?p=taxdetails%26id=597352, accessed 15 May 2017.
365. WoRMS, http://www.marinespecies.org/aphia.php?p=taxdetails%26id=558218, accessed 28 July 2017.
366. WoRMS, http://www.marinespecies.org/aphia.php?p=taxdetails%26id=597335, accessed 11^th May 2017.
367. WoRMS, http://www.marinespecies.org/aphia.php?p=taxdetails%26id=597336, accessed 15 May 2017.
368. WoRMS, http://www.marinespecies.org/aphia.php?p=taxdetails%26id=597358, accessed 11^th May 2017.
369. WoRMS, http://www.marinespecies.org/aphia.php?p=taxdetails%26id=558319, accessed 28 July 2017.
370. WoRMS, http://www.marinespecies.org/aphia.php?p=taxdetails%26id=597339, accessed 15 May 2017.
371. WoRMS, http://www.marinespecies.org/aphia.php?p=taxdetails%26id=597403, accessed 15 May 2017.
372. WoRMS, http://www.marinespecies.org/aphia.php?p=taxdetails%26id=597451, accessed 15 May 2017.
373. WoRMS, http://www.marinespecies.org/aphia.php?p=taxdetails%26id=209601, accessed 28 July 2017.
374. WoRMS, http://www.marinespecies.org/aphia.php?p=taxdetails%26id=597452, accessed 15 May 2017.
375. WoRMS, http://www.marinespecies.org/aphia.php?p=taxdetails%26id=597406, accessed 15 May 2017.
376. WoRMS, http://www.marinespecies.org/aphia.php?p=taxdetails%26id=597365, accessed 14 June 2017.
377. WoRMS, http://www.marinespecies.org/aphia.php?p=taxdetails%26id=597367, accessed 28 July 2017.
378. WoRMS, http://www.marinespecies.org/aphia.php?p=taxdetails%26id=597341, accessed 15 May 2017.
379. WoRMS, http://www.marinespecies.org/aphia.php?p=taxdetails%26id=597380, accessed 11 May, 2017.
380. WoRMS, http://www.marinespecies.org/aphia.php?p=taxdetails%26id=597343, accessed 15 May 2017.
381. WoRMS, http://www.marinespecies.org/aphia.php?p=taxdetails%26id=597374, accessed 14 June 2017.
382. WoRMS, http://www.marinespecies.org/aphia.php?p=taxdetails%26id=559960, accessed 15 May 2017.
383. WoRMS, http://www.marinespecies.org/aphia.php?p=taxdetails%26id=884441, accessed 28 July, 2017.
384. WoRMS, http://www.marinespecies.org/aphia.php?p=taxdetails%26id=597315, accessed 11 May 2017.
385. WoRMS, http://www.marinespecies.org/aphia.php?p=taxdetails%26id=597398, accessed 11 May, 2017.
386. WoRMS, http://www.marinespecies.org/aphia.php?p=taxdetails%26id=139143, accessed 11 May, 2017.
387. WoRMS, http://www.marinespecies.org/aphia.php?p=taxdetails%26id=597530, accessed 11 May, 2017.
388. WoRMS, http://www.marinespecies.org/aphia.php?p=taxdetails%26id=597522, accessed 11 May, 2017.
389. WoRMS, http://www.marinespecies.org/aphia.php?p=taxdetails%26id=597514, accessed 15 May 2017.
390. WoRMS, http://www.marinespecies.org/aphia.php?p=taxdetails%26id=597520, accessed 15 May 2017.
391. WoRMS, http://www.marinespecies.org/aphia.php?p=taxdetails%26id=597543, accessed 15 May 2017.
392. WoRMS, http://www.marinespecies.org/aphia.php?p=taxdetails%26id=558680, accessed 21 September 2017.
393. WoRMS, http://www.marinespecies.org/aphia.php?p=taxdetails%26id=597820, accessed 15 May 2017.
394. WoRMS, http://www.marinespecies.org/aphia.php?p=taxdetails%26id=597823, accessed 15 May 2017.
395. WoRMS, http://www.marinespecies.org/aphia.php?p=taxdetails%26id=558649, accessed 15 May 2017.
396. WoRMS, http://www.marinespecies.org/aphia.php?p=taxdetails%26id=597827, accessed 15 May 2017.
397. WoRMS, http://www.marinespecies.org/aphia.php?p=taxdetails%26id=597533, accessed 15 May 2017.
398. WoRMS, http://www.marinespecies.org/aphia.php?p=taxdetails%26id=597534, accessed 15 May 2017.
399. WoRMS, http://www.marinespecies.org/aphia.php?p=taxdetails%26id=597536, accessed 15 May 2017.
400. WoRMS, http://www.marinespecies.org/aphia.php?p=taxdetails%26id=597522, accessed 15 May 2017.
401. WoRMS, http://www.marinespecies.org/aphia.php?p=taxdetails%26id=597538, accessed 15 May 2017.
402. WoRMS, http://www.marinespecies.org/aphia.php?p=taxdetails%26id=558646, accessed 28 July 2017.
403. WoRMS, http://www.marinespecies.org/aphia.php?p=taxdetails%26id=536910, accessed 28 July 2017.
404. WoRMS, http://www.marinespecies.org/aphia.php?p=taxdetails%26id=594422, accessed 9^th May 2017.
405. WoRMS, http://www.marinespecies.org/aphia.php?p=taxdetails%26id=536827, accessed 9^th May 2017.
406. WoRMS, http://www.marinespecies.org/aphia.php?p=taxdetails%26id=872299, accessed 9^th May 2017.
407. WoRMS, http://www.marinespecies.org/aphia.php?p=taxdetails%26id=581816, accessed 9^th May 2017.
408. WoRMS, http://www.marinespecies.org/aphia.php?p=taxdetails%26id=581817, accessed 15 May 2017.
409. WoRMS, http://www.marinespecies.org/aphia.php?p=taxdetails%26id=181228, accessed 15 May 2017.
410. WoRMS, http://www.marinespecies.org/aphia.php?p=taxdetails%26id=411132, accessed 15 May 2017.
411. WoRMS, http://www.marinespecies.org/aphia.php?p=taxdetails%26id=599485, accessed 23 May 2017.
412. WoRMS, http://www.marinespecies.org/aphia.php?p=taxdetails%26id=536695, accessed 9^th May 2017.
413. WoRMS, http://www.marinespecies.org/aphia.php?p=taxdetails%26id=536574, accessed 9^th May 2017.
414. WoRMS, http://www.marinespecies.org/aphia.php?p=taxdetails%26id=531260, accessed 30 May 2017.
415. WoRMS, http://www.marinespecies.org/aphia.php?p=taxdetails%26id=730410, accessed 28 April 2017.
416. WoRMS, http://www.marinespecies.org/aphia.php?p=taxdetails%26id=456805, accessed 28 April 2017.
417. WoRMS, http://www.marinespecies.org/aphia.php?p=taxdetails%26id=457000, accessed 1 June 2017.
418. WoRMS, http://www.marinespecies.org/aphia.php?p=taxdetails%26id=204424, accessed 28 April 2017.
419. WoRMS, http://www.marinespecies.org/aphia.php?p=taxdetails%26id=890630, accessed 21 September 2017.
420. WoRMS, http://www.marinespecies.org/aphia.php?p=taxdetails%26id=412595, accessed 1 June 2017.
421. WoRMS, http://www.marinespecies.org/aphia.php?p=taxdetails%26id=558400, accessed 1 June 2017.
422. WoRMS, http://www.marinespecies.org/aphia.php?p=taxdetails%26id=851422, accessed 28 July 2017.
423. WoRMS, http://www.marinespecies.org/aphia.php?p=taxdetails%26id=572093, accessed 27 April 2017.

---