DEEP-SEA CORAL COLLECTION PROTOCOLS

A synthesis of field experience from deep-sea coral researchers,

designed to build our national capacity to document deep-sea coral diversity

Etnoyer, P., S. D. Cairns, J. A. Sanchez,

J. K. Reed, J.V. Lopez, W.W. Schroeder,

S. D. Brooke, L. Watling, A. Baco-Taylor,

G. C. Williams, A. Lindner, S. C. France,

and A.W. Bruckner

U.S. Department of Commerce

National Oceanic and Atmospheric Administration

National Marine Fisheries Service

NOAA Technical Memorandum NMFS-OPR-28

August 2006
Citation for the document:

Etnoyer, P., S. D. Cairns, J. A. Sanchez, J. K. Reed, J.V. Lopez, W.W. Schroeder, S. D. Brooke,

L. Watling, A. Baco-Taylor, G. C. Williams, A. Lindner, S. C. France, and A.W. Bruckner. 2006.

Deep-Sea Coral Collection Protocols. NOAA Technical Memorandum NMFS-OPR-28, Silver

Spring, MD. 53 pp.

For more information:

Additional copies of this publication can be obtained from: Andrew Bruckner, NOAA Fisheries, Office

of Habitat Conservation, Ecosystem Assessment Division, 1315 East West Highway, Silver Spring,

MD 20910. 301-713-3459.

i

Acknowledgements

Special thanks to NOAA’s Office of Ocean Exploration for supporting this work, to Liam Antrim for

compiling information on shipping deep-sea coral material. Lance Morgan and Elliott Norse of the

Marine Conservation Biology Institute helped to launch the project early on. Margaret Pizer and Fan

Tsao provided editing assistance. Several NOAA personnel provided valuable insights and

encouragement to this document, notably John McDonough, Andrew Shepard, Kim Pugliese, and Brad

Barr. A mini-grant from The Census of Marine Life project CenSeam (a global census of marine life on

seamounts) helped to bring this project to completion. The authors are grateful for constructive

comments provided by Frank Parish, Bob Stone, Nicolas Alvarado and two other anonymous

reviewers. Funding for printing was provided by NOAA Fisheries, Office of Habitat Conservation.

Tom Hourigan and Andy Bruckner from NOAA’s Office of Habitat Conservation brought this

document to press.

DISCLAIMER:

This publication does not constitute an endorsement of any commercial product or intend to be an

opinion beyond scientific methodologies and results compiled by the authors. The opinions and views

expressed in this document are those of the authors and do not necessarily reflect those of the National

Marine Fisheries Service, National Oceanic and Atmospheric Administration, or the U.S. Government.
U.S. Department of Commerce

Carlos Gutierrez, Secretary

National Oceanic and Atmospheric Administration

Vice Admiral Conrad C. Lautenbacher, Jr., USN (Ret.)

Under Secretary for Oceans and Atmosphere

National Marine Fisheries Service

William T. Hogarth, Assistant Administrator for Fisheries

NOAA Technical Memorandum NMFS-OPR-28

August 2006

DEEP-SEA CORAL COLLECTION PROTOCOLS

A synthesis of field experience from deep-sea coral researchers,

designed to build our national capacity to document deep-sea coral diversity

Etnoyer, P., S. D. Cairns, J. A. Sanchez, J. K. Reed, J.V. Lopez, W.W. Schroeder, S. D. Brooke,

L. Watling, A. Baco-Taylor, G. C. Williams, A. Lindner, S. C. France, and A.W. Bruckner

ii

Office of Habitat Conservation, Rm 15829

NOAA National Marine Fisheries Service

1315 East-West Highway

Silver Spring, MD 20910-3282

The sea pen Pteroeides sp., 9-320 m
iii

Around the time that the thirteen original Atlantic colonies were fighting for independence from

Britain, there existed little agreement among naturalists as to the nature of corals. Were they inanimate

(stones), plants, animals, or intermediate between the latter two (zoophytes)? This diversity of definition

and opinions undoubtedly produced considerable confusion and disagreement among naturalists

interested in such things. The symbiotic nature of algal cells in the tissues of some corals was also not

well understood. It was not until the Darwinian period in the nineteenth century that little doubt

remained, and therefore it was generally agreed, that corals were actually animals – heterotrophic living

organisms that prey on other organisms for nutrition and do not produce their own food.

In the past fifty years the basic goals and tenets of deep-sea coral collection, curation, and

taxonomy have changed little. On the other hand, the techniques and tools of this particular avenue of

research have changed significantly. Regarding the collection of material in the field, some aspects

remain fundamentally the same. The use of research vessels, bottom trawls, and naturalist’s dredges are

still frequently used for deep water research. In shallow water collecting, improvements in SCUBA

diving equipment and new innovations, such as Trimix gas and Nitrox diving, have allowed divers to

work at greater depths with longer bottom times. Pressure independent dive suits have permitted

researchers to attain depths not possible in traditional wet or dry suits. In the past four decades,

advances in optics, electronics, and robotic technology have allowed for a rapid sophistication and a

broader scope of possibilities regarding manned submersibles, remotely operated vehicles (ROV’s), and

more recently, autonomous underwater vehicles (AUV’s). Great strides have been made since the early

1990’s in the technological aspects of the collection and photography of the deep water benthos.

Concerning the techniques and tools of research in the laboratory, the scanning electron

microscope has for several decades provided a valuable tool for documenting surface details of

sclerites. Newer electron microscopes and digital technology have negated the necessity of using

photographic film and images can now be unloaded directly on to computers for processing. Relatively

recently, the fields of phylogenetics, molecular biology, and natural products biochemistry have been

applied to more traditional modes of research in the field of octocoral systematics.

This document on deep-sea coral collection protocols stands as a valuable resource, because it

provides a standardized procedure for researchers during a time of rapidly changing technology

regarding exploration of the deep-sea benthos. Stated another war, the protocols give us an instruction

manual for research procedures in deep-sea benthic fieldwork. Technological improvements coupled

with workable field-tested procedures (such as provided by the present volume) can in the long run,

only help to expand and improve our base of knowledge concerning octocoral diversity.

The synthesis of results from research endeavors both old and new, and opportunities allowed

by a constantly improving technological scene, provide for a positive potential for the future in the fields

of octocoral diversity and systematics.

Forward

Gary C. Williams, California Academy of Sciences
 iv

Table of Contents

Forward ......................................................................................... iii

Introduction..................................................................................... 1

Deep coral habitat and range................................................................ 3

Notes on deep-sea coral biology

Branching patterns in octocorals................................................... 5

Sclerites in octocorals ............................................................... 9

Ahermatypic scleractinians of the western Atlantic ............................ 11

Reproduction in deep-sea colonial corals ........................................ 13

Genetic methods for species identification, evolution, and ecology ......... 17

Handling procedures adopted by the Mountains-in-the-Sea Research

Group ................................................................................. 19

Documenting deep-sea coral collections

Video protocol ...................................................................... 23

Transect types ...................................................................... 25

Still image protocol ................................................................ 26

Living Specimen Photography .................................................... 27

Collecting .................................................................................... 29

Permits ....................................................................................... 29

Preserving .................................................................................... 30

DSC preservation methods tear sheet ......................................... 34

Labeling ....................................................................................... 35

Shipping ....................................................................................... 36

References and further reading .......................................................... 37

Image credits ............................................................................... 44

Appendix 1. Classification of the Deep-sea Corals .................................... 45

Appendix 2. Deep-sea coral Panel of Experts ........................................... 47

Appendix 3. Deep-sea corals Online...................................................... 49
1

what we know is garnered from early oceanographic expeditions that dragged the seafloor with trawl

nets. Dried broken fragments fill museum drawers and scientific literature, but living coral communities

from the deep-sea still remain largely unexplored.

Things are changing rapidly. Worldwide, ocean exploration activity is increasing. Interest in deep-sea

coral and seamount habitats is also increasing. New technologies are taking us deeper, longer, with

better tools. Around the globe, researchers are discovering rich benthic communities, and abundant

habitat. As a result, pressure is growing on scientific institutions to curate and identify deep corals and

other invertebrates. The scientific community is poised for new understanding, using new sampling

techniques, and new database technologies that associate myriad data types with physical samples;

including maps, still and moving images, and DNA barcodes. New standards will be necessary. Marine

invertebrate sample collection protocols for deep-sea organisms are well known within some institutions

but, until now, there were no publicly available, multi-institutional, peer-reviewed methodologies

describing how to collect deep-sea coral samples to meet archival standards.

NOAA’s Office of Ocean Exploration commissioned these collection protocols to facilitate greater

understanding, and to increase our national capacity to document deep-coral diversity in federal waters.

This document is designed to partly to provide useful background information, and partly to build

consensus among researchers, but mostly to guide collections by non-specialists and non-biologists.

Marine archeologists, geologists, fishermen, oilmen, and divers can all make useful scientific collections

given proper tools and techniques, so we try to share those techniques here. Each aspect described

herein will not always be possible or necessary, so we endeavor to provide simple collection methods

that will bring useful biological specimens to a broad spectrum of scientific disciplines.

In reading, you will encounter a variety of techniques and approaches from field scientists across the

country in a variety of disciplines. Consensus was not entirely possible within this group, due to the

different needs of different disciplines, so we list alternatives wherever we can. The reader must choose

what works best for their goals, with equipment at hand. Collectively, the assembled authors provide

good background information on deep coral biology, morphology, and habitat. The document is unique

for its attention to octocorals. The authors cover important technical aspects of coral collection in some

Fig. 1. Septa of cup coral, Desmophyllum sp.

Introduction

Most of our knowledge about corals comes from shallow

water tropical reefs and laboratories. These studies

showed us that corals are diverse, delicate, slow-growing,

and long-lived. They showed us that reefs are vulnerable

to nutrient input, climate change, and destructive fishing

gear, but healthy coral reefs can protect coastal

communities, and provide habitat for numerous associated

species. Yet, corals are not restricted to tropical reefs.

Corals grow in many types of seafloor habitats ranging

from the shallows to the abyss, and from the equator to

the poles (Williams, 1990). Not surprisingly, corals from

“the other 90%” of the ocean, the deep-sea (>200m),are

still very poorly known and poorly understood. Most of
2

What is a deep-sea coral?

In this document, we refer to deep-sea corals as a loosely defined paraphyletic assemblage of

hexacoral, octocoral, and calcified hydroid families in the phylum Cnidaria with species known to grow

deeper than 200m, beyond the boundaries commonly attributed to zooxanthellate shallow water

hermatypic scleractinian corals. These are also called cold-water corals and deep-water corals (Cairns,

in press). Fishermen call gorgonians “trees”, and ahermatypic corals “spiders”. The majority of the

families described in this document are octocorals (Anthozoa: Alcyonaria: Alcyonacea) but some are

hexacorals (Anthozoa: Zoantharia: Antipatharia and Scleractinia).

Deep-sea corals come in many forms, ranging in size from 2 mm rice grain-sized solitary corals, to fistsized soft colonies, feathery sea pens, bushy black corals, 2 m long whip corals, large flabellate sea fans

(gorgonians), and massive bank framework scleractinians. Deep-sea corals are diverse and beautiful,

and they range over of a greater part of the coastal United States than the more familiar zooxanthellate

tropical corals. Many species have complex branching morphologies and grow large enough to form

structure for associated species. Other species, like sea pens, have simple pinnate forms but they grow

in abundance, and provide important ecological habitat in an otherwise flat and featureless abyss. Some

coral colonies are long-lived. They function as archives of climate change, and they are uniquely adapted

to extreme environments. Deep-sea corals are an exciting and fast growing field of research, and an

integral part of NOAA’s agenda for deep-sea exploration.

A complete taxonomic list of 48 families of hexacorals, octocorals, and calcified hydroids with species

known to occur deeper than 200m is provided in Appendix 1- Classification of Deep-sea Corals.

This list describes the families for which these protocols will be most useful.

Fig. 2. Bamboo coral (Isidella sp.) polyps. Fig. 3. Living bamboo coral (Isidella sp.) colony

detail: including documenting your specimen in-situ, collecting your specimen to insure scientific

value, preserving your specimen to enhance long-term study, labeling your specimen to insure

accurate transcription, and shipping your specimen to accredited scientific institutions. We hope you

find the information useful.
3

Depth and frequency of occurrence for selected 

deep-sea coral families 

0%

10%

20%

30%

40%

50%

60%

70%

0 400 800 1200 1600 2000 2400 2800

Depth (m) 

Frequency of occurrences

Primnoidae (n=925)

Paragorgiidae (n=257)

Isididae (n=335)

Antipathidae (n=208)

Fig. 4. Depth and frequency for some Northeast Pacific

octocoral families (from Etnoyer and Morgan 2005).

Deep-sea coral habitat and range

by Peter Etnoyer

When most people think of coral reefs, they

think of shallow water tropical reefs. However,

deep coral communities of some type lie off the

shore of most coastal nations and states, as

deep as 6000m. Contrary to popular opinion,

both the hard scleractinian hexacorals (the

zoantharia) and softer gorgonaceous octocorals

(the alcyonaria) have vertical ranges exceeding

6000m. They range worldwide. The higher level

taxa (families) are generally cosmopolitan, with

some worlwide genera, and some strictly

regional genera.

Perhaps the deepest known coral taxon is a sea

pen — genus Umbellula, from 6620m

 (Williams, 1999). Primnoids occur as deep as 5850m (S. Cairns, pers. comm.) and thrive in

abundance as deep as 3000m (A. Baco-Taylor, pers. comm.). The deepest known bamboo coral

Keratoisis profunda, at 4851m (Cimberg, 1981) near the Northwest Hawaiian Islands. Hydrozoan

‘lace corals’ have a worldwide distribution but are most common near islands and seamounts. Lace

corals have a vertical range from 0.5 to 2,787m (Cairns, 1992). Ahermatypic ‘thickets’ of Lophelia

pertusa in the Gulf and the US South Atlantic bight are generally found between 200 and 1000m

(Schroeder et al. 2005).) Most of the coelenterate families that contain deep-sea corals include shallowwater species, too.

Deep-sea corals can occur in many assemblages, including monotypic scleractinian bioherms (e.g.

Lophelia sp. in the Atlantic), monotypic octocoral fields or meadows in mixed substrates on low relief

mounds (e.g. Callogorgia and Acanella in the Gulf of Mexico) or diverse rocky reefs of octocorals,

anemones and sponges (e.g. New England and Gulf of Alaska Seamounts, Aleutian and Hawaiian

Fig. 5. Stony corals and whip corals on hardbottom habitat.

Islands). Beginning in the North Atlantic

and traveling clockwise around the

country, we find Paragorgia sp.

bubblegum corals are common to

1600m on the New England seamounts

(L. Watling, pers. comm.). Rare and

fascinating octocorals like the beautiful

Metallogorgia and Iridogorgia bright

stony corals like Enallopsammia

pictured here are also found there

(Watling and Auster, 2005). South
4

Atlantic Bight is known for some large banks of Oculina varicosa and lithoherms of L. pertusa that

provide habitat for more than 58 species of fish, and almost 142 species of benthic invertebrates (Reed

et al., 2006). Some of these lithoherms have been impacted by destructive fishing gear, but are now

coming under protection. The Gulf of Mexico is known for deep thickets of L. pertusa (Schroeder et al.

2002), though several species of octocorals occur there. L. pertusa colonies can encrust much of a

sixty-year-old shipwreck (at 550m) with the largest thickets 6m tall and 3.5m wide (W. Schroeder, pers

comm.). Images and videos from the shipwrecks can be viewed at the Past Foundation website:

http://www.pastfoundation.org/DeepWrecks/Video03.htm. L. pertusa also encrusts oil rigs.

In the North Pacific, colorful hydrocoral banks like the Stylaster mound pictured on the cover are found

25m and deeper as far south as Baja California. These banks are well photographed, and very popular

with sport divers and fishermen. Large black, bubblegum, primnoid, and bamboo corals across can be

found deeper than 100m on the continental shelves of California, Oregon and Washington states. Coral

forests in these regions are thought to play important roles as habitat for Atka mackerel and rockfish

(Heifetz 2002, Krieger and Wing 2002). Shallow seamount peaks in the Gulf of Alaska host similar

species. The Aleutian Islands of Alaska are incredibly rich. Submerged banks and peaks in the Hawaiian

islands are famous for black, gold, and precious corals once harvested for the jewelry trade (Grigg,

1976). Several species are now listed as endangered.

Some solitary cup corals (e.g. Desmophyllum) thrive on small ledges and overhangs, while others (e.g.

Balanophylla) settle on cobbles and pebbles. Gorgonians thrive in rocky habitats with strong currents.

Currents facilitate settlement onto clean swept surfaces, increase food availability and capture and,

therefore, growth rate and survivorship (Patterson 1984, Sponaugle and LaBarbera 1991). Octocorals

have been used as indicators of flow velocity on seamount peaks (Genin et al. 1986), orienting

perpendicular to prevailing currents (Grigg, 1972) but some species, like the Acanella bamboo corals

and the sea pens are also adapted to muddy and mixed substrates of low-relief (Smith et al 2002,

Williams 1995). Interestingly, small differences in relief (e.g. boulders ~1m) also provide deep coral

habitat in an otherwise featureless plain.

Fig. 6. Galatheid crab

nestled in outstretched

octocoral polyps
5

Branching patterns in octocorals

by Juan A. Sánchez

Octocorals are one of many organisms that grow producing multi-branched tree-like colonies.

Interestingly, recent molecular phylogenies of octocorals show how branching can evolve independently

multiple times from non-branching hypothetical ancestors involving different axial materials (Sánchez et

al., 2003). Consequently, a particular branching pattern should not be considered taxon- or taxaspecific. There are remarkable cases of convergent evolution (i.e., analogy without common ancestry)

such as nearly identical pinnate morphologies in multiple octocorals, as in the case of

Pseudopterogorgia (Gorgoniidae), Plumigorgia (Ifalukellidae), Pteronisis (Isididae), and Plumarella

(Primnoidae; see Fig. 7), which differ in their modular (e.g., polyp and calyx) and ultrastuctural (e.g.,

sclerites) nature. This suggests caution when using branching morphologies as diagnostic characteristics

but on the other hand calls to attention the remarkable plasticity in the evolution of colony form in

octocorals. Figure 8 shows a sample of the diverse branching patterns and other colonial morphologies

observed in octocorals.

Taxonomists recognize up to 28 terms

for the description of octocoral colony

shapes and branching patterns (Bayer et

al., 1983), which can be redundant or

related in some cases. In order to

understand the branching pattern of

octocorals it is important to know how

they grow and branch. Branching in most

octocorals is a subapical process (i.e.,

just below the tip) where some

branches, “mother branches,” produce

new branches, “daughter branches,” at

roughly fixed distances or internodes

(Sánchez et al., 2004).

The extreme example of this is pinnate pattern where a single mother branch retains multiple daughter

branches (Fig. 7). This morphology is a kind of monopodial branching, of which the opposite extreme

example is only one “daughter” allowed for every “mother branch” producing a sympodial branching

rarely found in octocorals (Chrysogorgia sp. in Figures 7h and 8b). This implies that the colony has

simple branch set points (i.e., one daughter branch at the time) whereas pinnate branching would have

double set points.

Simple branching seems to be the most common in octocorals, generating pseudo-dichotomous (e.g.,

sea candelabrum) and pectinate/lyrate (e.g., Ctenocella pectinata) morphologies all growing in one

plane or uniplanar. If the tips of daughter branches from uniplanar morphologies join, or anastomose,

that creates reticulate branching, examples of this weblike pattern are the common sea fans (e.g.,

Gorgonia, Pacifigorgia, Subergorgia, etc.).

Fig. 7. Examples of pinnate branching patterns in octocorals. Left:

Pteronisis incerta (from Alderslade, 1998) and Pseudopterogorgia

bipinnata (Colombia, Caribbean).
6

Fig. 8. A. Sarcophyton glaucum (Quo and Gaimard), Great Barrier Reef, Australia, Western Pacific, shallow water (United

States National Museum catalog number-USNM: 79445, at NMNH, Smithsonian Institution, Washington D.C., USA). B.

Alcyonium digitatum Linnaeus, off New Jersey, USA, Western Atlantic, 85 m in water depth (USNM 60336). C. Anthothela

nuttingi Bayer, Hawaii, North Pacific, 1010 m (USNM 94435). D. Lepidisis olapa Muzik, Hawaii, North Pacific, 400 m

(USNM 56717). E. Anthomuricea tenuispina Nutting, Hawai, North Pacific, 517 m (USNM 98801) F. Paramuricea

hawaiiensis Nutting, Hawaii, North Pacific, 345 m (USNM 58385). G. Muricea fruticosa Verrill, Panama, Eastern Pacific, 60 m

(USNM 57102) H. Chrysogorgia chryseis Bayer and Stefani, Hawaii, North Pacific, 1010 m (USNM 100848). I. Lophogorgia

alba Duchassaing and Michelotti, Panama, Eastern Pacific, intertidal (USNM 57091). J. Leptogorgia virgulata, Gulf of Mexico,

USA, Western Atlantic, 6 m (USNM 49690). K. Renilla reniformis (Pallas), Puerto Rico, Caribbean, 2 m (USNM 79448). L.

Corallium ducale Bayer, Hawaii, North Atlantic, 1205 m (USNM 94458). M. Narella dichotoma (Verluys), Hawaii, North

Atlantic, 1218 m (USNM 98822). N. Isidella elongata (Esper), Italy, Mediterranean, 350 m (USNM 57457). O. Paragorgia

dendroides Bayer, Hawaii, North Pacific, 1018 m (USNM 98789). P. Acanthogorgia sp. Hawaii, North Pacific (USNM

100340). Scale bar = 2 cm. (from Sanchez et al., 2003a).
7

Fig. 9. Examples of deep-water corals fom New Zealand (NIWA collection). A. Metallogorgia melanotrichos;

B. Chrysogorgia sp.; C. axial view of B.; D. Keratoisis sp. and branching set point detail; E. Iridogorgia branching

pattern and axial sample; F. Acanella sp. and close-up detail. G. Thouarella sp. (scale is a US Lincoln penny).
8

Other colonies can modify and twist their axes to branch in multiple planes such as the spectacular

Iridigorgia colonies, which have lateral simple branching in a spiralized axis (Fig. 9E). Other double

branching morphologies, besides pinnate, are alternate branched colonies, which generate pseudodichotomous morphologies in the simpler cases (e.g., uniplanar) or bottlebrush colonies in more

complicated spiral arrangements (e.g., Chrysogorgia: Fig. 9B).

Pinnate colonies can also produce branches in multiple directions or bottlebrush morphologies (e.g.,

Thouarella: Fig. 9G). In most cases, you will find that many species exhibit combined and more

disorganized branching patterns creating simple bush-like colonies (e.g., Acanella: Fig. 9F). Some

combined morphologies present remarkable shapes such as Metallogorgia melanotichos, where the

colony is a long whip-like mother branch that commences branching on multiple sympodial fronts (e.g.,

Fig. 9A). In summary, octocoral branching patterns are a complex combination and continuum of

features such as branching set point (simple lateral/alternate or pinnate), planes of branching (uniplanar,

multiplanar, spiral, bushy, etc.) and generational relationships of branches (single mother branch:

monopodial to multiple mother branches: sympodial).

It is important to point out certain considerations for getting complete branching information from

collections of deep-water octocorals. Undoubtedly, deep-water octocorals exhibit the most interesting

and elaborated branching morphologies (e.g., Fig. 9). Nonetheless, their sampling is more delicate and

samples are more difficult to preserve (e.g., Fig. 9E). For closely-related species distinctions, branching

patterns often offer important characters, information which should particularly be collected for

Chrysogorgiidae, Primnoidae and Isididae. Branching set points are highly important for Isididae where

branching from the node or internode could differentiate Keratoisis from other Keratoisidinae (e.g.,

Fig. 9D, F).

The orientation of the polyp armatures with respect to the branching can also be of importance in

Primnoidae, where some can be upward, outward or downward (e.g., downward, Narella sp.

Fig. 10. Narella sp. (New Zealand, NIWA collection) with arrows

showing downward orientation of polyp sclerite armature.

Fig. 10). If the material cannot be

collected in its entirety it is important

to take photographs and details of

branching points, nodes, daughter

branch detail, and of course the

whole colony. An ideal situation for

remotely operated vehicles would be

to take 3-D stereo-pairs of the

colonies at various distances.
9

Sclerites in octocorals

by Stephen D. Cairns

Although branching morphology, corallum size, and

color are valuable hints for the identification of

octocorals, and are often the only characters

available for field identifications, the single most

important character in octocoral identification is the

microscopic calcareous sclerite (Bayer, 1956).

Sclerites, also called “spicules”, are found in almost

all 2800 octocoral species. They usually occur in

great numbers in every colony, 10’s of thousands

not being unusual in a modest sized colony. Even a

small fragment may contain thousands of sclerites.

Each sclerite is a small piece of calcitic calcium

carbonate ranging in size from 20 um to 5 mm; they

are found in the polyps of the colony as well as in

the coenenchymal tissue between the polyps, each

giving a small amount of support and structure to the

tissue and polyps. Fig. 11a. An illustrated octocoral axis and polyps.

Sclerites occur in many shapes and sizes, the first attempt at classifying them was the illustrated trilingual

glossary edited by Bayer, Grasshoff, & Verseveldt (1983), which defined, synonymized, and illustrated

57 sclerite types, some with the whimsical, yet descriptive, names of: caterpillar, crutch, finger-biscuit,

hockey-stick, rooted head, wart club, capstan, and opera-glass. A species may have only one type and

size of sclerite or a variety, each type sclerite type occurring in a particular part of the colony and serving

a slightly different function.

It is easy to examine the sclerite types from a colony. One simply cuts a small piece containing a polyp

and branch from a larger colony, place it on a microscope glass slide, and add a drop of common

household bleach. Wait a few minutes, or until the bubbles burst, place a cover slide over the

preparation (optional), and examine at x200 with a compound microscope, or as little as x50-100 with

a dissecting microscope. SEM of sclerites done at x400-500 or higher magnifications is usually not

necessary for routine identifications but does provide excellent illustrations that can be used to describe

species.

Fig. 11b.
10

For cleaner and more permanent preparations one should rinse the sample with successive solutions of

hydrogen peroxide, tap water, distilled water, 70% ethanol, and 95% ethanol (see Fabricius &

Alderslade, 2001: 38-39). It is important not to preserve octocorals in formalin, because sclerite

dissolution will occur over a period of several weeks to months. Specimens may be fixed for a short

time, (less than 24 hours) in a weak (~ 4%) formalin solution that is highly buffered.

The ability to identify the suite of sclerites of a specimen using Bayer, Grasshoff & Verseveldt (1983)

and using the key to the genera (Bayer, 1981) is an excellent start to the identification of any octocoral,

exclusive of pennatulids. As knowledge of sclerites is so important for the identification of octocorals, it

is not recommended to ask an octocoral specialist to identify a specimen based on a picture or video,

because his invariable response will be to ask for the specimen, if only a tiny fragment containing a few

sclerites.

135. Spiny ball. Asterospicularia randalli Gawel. x625.

136. Unilaterally spiny spheroid Mopsella sp. x250.

137. Triradiate. Titanideum ftauenfeldii Kolliker. x350.

138. Cross. Cavernulina sp. x300.

139. Cross. Ellisella sp. x375.

140. Cross. Gersemia rubiformis Ehrenberg. x375.

141. Six radiate. Corallium borneense Bayer. x375.

142, Six radiate. Paragorgia arborea Linnaeus. x750.

143. Six radiate. Epiphaxum micropora Bayer & Muzik. x375

144. Six radiate. Alcyonium digitatum Linnaeus. x500.

Fig. 11c.

Fig 11b-11c. Types of sclerites found in octocorals:

Figures 11a-c are reporduced from the Illustrated Trilingual Glossary of Morphological and

Anatomical Terms Applied to Octocorallia edited by Frederick Bayer, Manfred Grasshoff, and

Jakob Verseveldt and published by E.J. Brill, The Netherlands, 1983.
11

Ahermatypic scleractinians of the western Atlantic

by John K. Reed

Western Atlantic Deep-Water Coral Reefs

The dominant corals forming deep-water reefs in the western Atlantic region off southeastern US and Gulf of

Mexico are Oculina varicosa, Lophelia pertusa, and Enallopsammia profunda (Reed 2002 a,b,

Schroeder et al. 2005). Other branching colonial scleractinia also occur, including Solenosmilia variabilis,

Madrepora oculata, and Madracis myriaster, as well as numerous solitary coral species (Cairns, 1979).

The deep-water Oculina bioherms form an extensive reef system at depths of 70-100 m along the shelf edge

and are only known off central eastern Florida. These reefs are comprised of numerous pinnacles and ridges,

3 to 35 m in height. In comparison, deep-water reefs of Lophelia pertusa and Enallopsammia profunda

corals often occur together at depths of 500-850 m from Florida to North Carolina and the Gulf of Mexico.

These form reef structures from 10 to 150 m in height. Deep-water Lophelia reefs also occur worldwide

and extensively off Norway and Scotland. The structure of both the Oculina and Lophelia reefs are often

similar. Each pinnacle may consist of a bank of unconsolidated sediment and coral debris that is capped on

the slopes and crest with living and dead colonies of coral.

Fig. 12. Colony and branch tip for : top - Oculina varicosa (80m); middle - Lophelia pertusa (490 m); bottom - Enallopsammia

 profunda (585 m). Scale lines = 1 cm; top left fig. scale = 5 cm. From Reed, J. K. 2002.

Fig. 13. Morphological features of coral corallite. From Cairns, S. 1981.

Fig. 14. Septa patterns of a coral calice. From Cairns, S. 1981.

12. 13.

14.
12

Scleractinian Coral Descriptions

Oculina varicosa Lesueur, 1820: In deep water (>60 m), O. varicosa forms spherical, dendroid, bushy

colonies that grow to 1 to 2 m in diameter and height (Fig. 12, top). Individual corals may coalesce forming

linear colonies 3-4 m in length or massive thickets of contiguous colonies on the slopes and crests of the deepwater reefs (Reed, 1980; Reed 2002 a,b). The branches average 3-6 mm in diameter and frequently

anastomose. Corallites are distributed spirally around the branches, and calices are generally 2-3 mm in

diameter with 3 cycles (24) of septa which extend as costae or ridges on the outside of the calices. The

deep-water form lacks zooxanthellae, whereas in shallow water O. varicosa is usually golden brown with

the algal symbiont. The shallow-water O. varicosa colonies average <30 cm in diameter,with thicker

branches, but do not form thickets or coral banks like the deep-water form. O. varicosa ranges from the

Caribbean to Bermuda at depths of 2-152 m (Verrill, 1902; Smith, 1971; Reed, 1980). The deep-water

Oculina reefs , however, are only known from 27o 32' N and 79o 59' W to 28o 59' N and 80o 07' W, and

at depths of 70-100 m.

Lophelia pertusa (Linnaeus, 1758) (= L. prolifera): Similar in gross morphology to Oculina, this coral also

forms massive, dendroid, bushy colonies, 10-150 cm in diameter, with anastomosing branches (Fig. 12,

middle). Branch tips are slender whereas the base may be several centimeters in diameter. Terminal

branches have opposite, alternately arranged corallites which may be variable in shape from round to elliptical

and somewhat flared or flower-like. Calices may be up to 15 mm in diameter and number of septa is variable

with 7 to 11 primary septa which generally extend beyond the lip of the cup as costae. The columella, at the

bottom of the corallite cup, is rare but may consist of 1 to 3 small rods. Its distribution ranges in the western

Atlantic from Nova Scotia to Brazil and the Gulf of Mexico, and also in the eastern Atlantic, Mediterranean,

Indian, and eastern Pacific Oceans at depths of 60-2170 m (Cairns, 1979).

Enallopsammia profunda (Pourtalès, 1867) (=Dendrophyllia profunda): This species also forms

dendroid, massive colonies up to 1 m in diameter (Fig. 12, bottom). Branches are about 1 cm in diameter

and less at the tips. Characteristic of this species are the prominent cone-shaped corallites which are 3-4 mm

in diameter and generally arranged alternately on opposite sides of the branches. There are generally 3 cycles

of 24 septa that do not extend beyond the rim of the cup. A small columella occurs at the base of the corallite

as a spongy mass. It is endemic to the western Atlantic and ranges from the Antilles in the Caribbean to

Massachusetts at depths of 146-1748 m (Cairns, 1979).

These three common deep-water reef species may be distinguished by the characteristics described

above. They also may be differentiated from other bushy colonial species that may co-occur on

western Atlantic deep reefs. These include Solensomilia variabilis, which occurs to 2165 m and is

most readily identified by its intratentacular budding. Thus the branch tips often consist of a pair of

calyces in various stages of splitting apart. Madracis myriaster occurs to 708 m and is distinguished by

only having 10 primary septa and usually a strong style projects from the center of the columella.

Madrepora carolina is another colonial that occurs in the Gulf of Mexico. Fig. 13 and 14 illustrate

features of the corallite, and septal patterns of a coral callice.
13

Reproduction in deep-sea colonial corals

by Sandra Brooke

As coastal species become more depleted, increased effort will be displaced into deep-sea fisheries,

which pose the greatest threat to deep-sea coral habitats. Recovery of a damaged ecosystem is

dependent on the ability of the keystone species to re-colonize affected areas. Corals can do this via

asexual fragmentation, which can clearly be seen in deepwater coral habitats, but the effectiveness of

this mechanism is limited, because fragments cannot travel far from the parent colony. Although colony

growth and fragmentation can compensate for local small-scale damage, re-colonization of large areas

requires larval recruitment, which is determined by larval lifespan and behavior, local currents, and postlarval survival. Knowledge of reproductive strategy, fecundity, and larval biology will enable us to assess

the potential of deep-sea corals to recover from natural and anthropogenic impacts.

the spermatogonia then differentiate into primary spermatocytes, which undergo meiosis to form

secondary spermatocytes then spermatids (Delvoye 1982). Mature gametes are released through the

mouth, (spawning cues are unknown), and embryogenesis and larval development take place in the

water column.

A study of reproduction in deep-sea corals from New Zealand showed that four colonial species,

Enallopsammia rostrata, Goniocorella dumosa, Madrepora oculata and Solenosmilia variabilis

were all gonochoristic: each colony was either male or female. Broadcast spawning was the probable

mode of reproduction, with spawning inferred in all species for late April or May. There was a high level

of synchrony between species in the seamount localities studied. High fecundities were estimated for E.

rostrata (>144 oocytes per polyp), G. dumosa (>480 oocytes per polyp) and S. variabilis (>290

oocytes per polyp), with a negative correlation between oocyte size and fecundity for all three species

(S. Burgess, pers .comm.).

Oculina varicosa is a gonochoristic broadcast-spawning species with an annual reproductive cycle.

Gametogenesis begins in early spring and terminates in a protracted spawning season that lasts from late

July until early September. Fecundity level varies from ~2000-5000 oocytes per cm2

 of skeletal surface

area. There is evidence for male-induced spawning in females, but again, information regarding

Fig. 15. Living polyps of Lophelia pertusa

Scleractinians do not have true reproductive organs;

eggs and sperm develop within the acellular mesogleal

layer of the mesenteries (Fig. 16). Gamete development

begins when primordial germ cells, appear within the

mesenterial gastrodermis (Szmant-Froelich et al. 1980,

1985, Delvoye 1982, Wyers 1985). These cells

proliferate and migrate from the gastrodermis into the

mesogloea of the mesenteries, where they undergo

meiosis to form primary oocytes. As oocyte

development proceeds, yolk is laid down and the

cytoplasmic volume increases (Fig. 16). Male germ

cells originate in the same way as females. Primordial

germ cells enter the mesogloea from the gastrodermis,
14

spawning patterns and cues is limited as spawning was only observed in the laboratory. Larvae are very

temperature tolerant and probably last between 14 and 21 days in the water column, which infers a high

dispersal potential. Larval biology has not been studied for any deepwater coral except O. varicosa

(Brooke and Young 2003).

The gametogenic cycle of Lophelia pertusa has been studied in samples from various locations; the

northeast Atlantic (Waller and Tyler in press), Norwegian Fjords (Brooke and Jarnegren in prep) and

the Gulf of Mexico (Brooke in prep). Lophelia pertusa is a pseudo colonial species, but has similar

reproductive traits (gonochoristic broadcast spawner) to the deep-water colonial species. In the North

Atlantic, the maximum oocyte diameter was 140ìm with a fecundity of 3146 oocytes per polyp.

Gametogenesis began in the summer (June-August) and spawning was inferred from histological

samples to occur between January and February. Samples from the Trondhjeim Fjord in Norway

showed a similar pattern of gametogenesis and spawning to those in the Northeast Atlantic, with

spawning probably occurring in February when the maximum average oocyte diameter (106ìm) was

observed. The samples from the Gulf of Mexico however show a completely different gametogenic

schedule. Vitellogenic oocytes (average diameter 80ìm) were observed in July, and samples collected

in early October show no signs of any gametogenic material, indicating that spawning occurs sometime

in late summer. The factors driving deepwater coral reproduction are still unknown.

Common traits can be observed in all the species studied: they are all gonochoristic broadcast spawning

species that reproduce seasonally. Deep-water corals live at depths beyond the photic zone (except for

O. varicosa, which may experience low light levels during the summer months) and rely exclusively on

planktonic food. Reproduction therefore may be driven by seasonal changes in food availability or by

changes in temperature, tidal regime, or some other environmental parameter. This is an interesting

avenue for further investigation, particularly since the reproductive cycles of L. pertusa are different in

samples from different locations.

Fig. 16. Light microscope images of Oculina varicosa mesenteries showing A) late vitellogenic oocytes; B) late

 stage testes packed with spermatids.
15

Although the focus of this chapter is reproduction in

the deepwater scleractinia, members of the

gorgonacea, antipatharia and stylasterina also fall

under the definition of colonial corals, and all may

comprise significant components of deepwater coral

ecosystems. Apart from the stylasterina, these orders

all have the same basic bauplan, with simple polyps

that contain the gametogenic material.

A study of Antipathes fiordensis from New Zealand

(Parker et al 1997) showed that like many colonial

scleractinians, this black coral species is

gonochoristic, with an approximate 1:1 sex ratio and a

seasonal reproductive cycle resulting in broadcast

spawning of gametes. The gametes originate in the

endoderm (gastrodermis) of the mesenteries but,

Fig. 16c. Eggs at the base of a bamboo coral polyp

The order Stylasterina (phylum Cnidaria, class Hydrozoa) are a group of calcareous encrusting or

branching colonial species often referred to as hydrocorals. Unlike scleractinians, there is no information

in the literature indicating that the survival and reattachment of branchfragments (fragmentation) is a

mechanism of new colony formation in hydrocorals. The male and female reproductive structures or

Fig. 16d. Longitudinal section of Metallogorgia

 melanotrichos polyp with developing oocytes

unlike scleractinians, the gametes do not migrate to the mesoglea to continue development. An

investigation into gametogenic cycles of 6 common gorgonian species in Biscayne Bay Florida

(Fitzsimmons-Sosa et al. 2004) showed that all species were gonochoristic, five showed reproductive

seasonality and sex ratios were either 1:1 or 1:2 male: female. As with the hexacorals, gametes of these

octocorals also develop within the mesenteries of the polyps (Fig. 16c).

gonophores develop inepidermally-lined cavities called

ampullae, which are usually present as superficial

hemispheres, but occasionally they are completely

submerged (internal) in the coenosteum. They are

generally smaller and more numerous in male colonies.

Stylasterines are gonochoristic, with internal fertilization

and embryonic development. Planular larvae are released

via efferent ducts, which are usually seen as small irregular

pores on or near the ampullae. A study of reproduction in

Alaskan hydrocorals (Brooke 2004), showed a range of

developmental stages in all species studied. This implies

either continuous reproduction or an extended nonsynchronous developmental season. Reproductive

strategy has implications for habitat recovery potential; for

example, a species with a longer planktonic phase may be

more able to recover from large scale damage than one

with a very short larval phase, or those that can form new

colonies from fragmented branches may recover quickly

from small scale physical damage.
16

To fully describe gametogenic cycles, samples should be collected monthly over multiple years.

However, the logistics and expense of working in the deep sea often prohibit such rigorous sampling

regimes. Small fragments (10-20 polyps) should be taken from several different colonies, preferably at

least 10. In a gonochoristic species with equal sex ratios, this will provide approximately 5 replicate

females, which is usually sufficient for statistical analysis, but more samples would give better statistical

power if logistics permit. Males can be used to describe stages of gonad development, but female

oocyte size frequency distribution is easier to quantify and is generally used to describe the gametogenic

cycle. Samples should be fixed while alive and in good condition. Stressed corals often produce

mucous, which can inhibit fixation so ideally, a cold room would be available to allow the corals to

recover from the stress of collection and exposure to warm surface water. If not, then fix the samples as

soon as they come on board. Dying or decaying tissue is not useful.

There are many different fixatives that can be used; the most readily available effective fixative to use is

10% formalin in seawater, which is made by diluting full-strength formaldehyde (37%) in a 1:10 ratio

with seawater. This is not recommended for fine cytological examination, but works sufficiently well to

determine gametogenic status. Tissues should be left in this fixative for at least 36 hours before

thoroughly rinsing in distilled water and transferring to70% ETOH for storage. Hellys solution (Barszcz

and Yevich 1975) is a better fixative for corals but is a little more complicated. A stock solution of 100

gm potassium dichromate and 200 gm zinc chloride in 4 L seawater can be made prior to sampling, but

the addition of the formaldehyde (200ml) should only be made when needed since the full mixture is

only good for 24 hours. The samples should remain in the solution for 16-20 hours then thoroughly

rinsed in freshwater and transferred to 70% ETOH for storage. Another alternative is ‘Z-fix’ (Anatech

Ltd.), which also contains formaldehyde, and is simply mixed with seawater prior to use.

Corals need to be decalcified before further processing. A solution of 10% hydrochloric acid with

EDTA buffer takes between 1 and 12 hours to decalcify, depending on the sample. An acid change may

be needed for heavily calcified skeletons. After rinsing thoroughly in running freshwater, tissues are

dehydrated through a series of ethanol concentrations (70-100%) and then transferred to a clearing

agent such as toluene. The tissues are then embedded in paraffin wax, cut into 8μm sections with a

microtome blade and stained using Mayer’s Haematoxylin/Eosin B before mounting on microscope

slides. This combination of stains will show lipids (vitellogenic oocytes) as bright pink and DNA

(primordial germ cells as testes) as purple. Oocyte diameter can be measured using a compound

microscope with an ocular micrometer or images of oocytes can taken using a digital camera and

subsequently measured with image analysis software. There will be some shrinkage of the oocyte during

the histological processing, so all measurements are “relative”. A minimum of 50 (preferably 100)

oocytes should be measured per polyp; however this is not always possible. This basic process can be

applied to any of the deepwater colonial corals, but for those species with large yolky eggs, paraffin

penetration can be improved by using a series of toluene and paraffin solutions.

A full treatise on histological techniques is beyond the scope of this chapter but recommended further

reading includes Carson, 1997, Kiernan, J.A. 2003.
17

Genetic methods for species identification, evolution, and ecology

by Amy R. Baco-Taylor

Genetic methods may be used to enhance taxonomic studies, to gain understanding of evolutionary

relationships, and to elucidate the dispersal patterns of deep-sea corals. Emerging molecular methods

may also be used to better understand deep-sea coral physiology. The key to molecular studies of any

taxon is to obtain informative markers, genes or segments of DNA that vary enough between individuals

or species to address the research hypotheses. Many informative phylogenetic and population genetic

markers have been developed for invertebrate taxa. Mitochondrial genes (particularly COI, 16S, cyt b,

and others) are commonly used. Unfortunately for coral biologists, many of these markers show little or

no variation in hexacorals or in octocorals (Van Oppen et al. 1999; France and Hoover 2002; reviewed

in Shearer and Coffroth 2004). Recent efforts have focused on new genes and microsatellite markers.

Some of these are reviewed below.

Fig. 17. Enallopsammia sp. on Manning Seamount.

phylogenetics, genes are sequenced in multiple individuals. and the resulting sequences are compared.

Differences in sequence composition are used to construct phylogenetic trees, which are used to

determine evolutionary relationships.

The primary markers used in phylogenetic studies of invertebrates are the nuclear gene ribosomal 18S

RNA and the mitochondrial genes ribosomal 16S RNA and cytochrome oxidase I (COI). 18S rRNA

has been used within the class Anthozoa (Berntson et al. 2001) and for higher-level systematics in

octocorals (Berntson et al. 1999; Sanchez et al. 2003a). Mitochondrial16S rRNA has also been used

to study systematics in shallow and deep-sea scleractinian corals (Romano and Palumbi 1996, 1997;

Romano and Cairns 2000) and in octocorals, (France et al. 1996). Mitochondrial 16S and COI have

low variability in scleractinian corals anbd octocorals compared to other invertebrates (reviewed in

Shearer et al. 2002). However, 16S has two insertion/deletion regions (INDELs, France and Hoover

2001). These INDEL regions have been used to separate species of bamboo corals (Smith et al. 2004)

and may be useful for higher-level coral systematics (Sanchez et al. 2003b).

Because Anthozoans have low variability in the genes typically used for phylogenetics, a number of other

genes have been tested in octocorals, including mitochondrial genes in the NADH group (ND2, ND3,

ND4, and ND6). Octocorals also have a mutS gene homolog in the mitochondrion. A portion of this

Phylogenetics and Systematics

DNA sequencing and phylogenetic analyses enhance

taxonomic studies based on morphology.

Phylogenetic analyses may be used to determine

whether two specimens are the same species, or

different species. They may be used to identify

cryptic species—species that are nearly

indistinguishable based on morphology but show

clear differences in their DNA sequences.

Phylogenetic analyses also help us understand the

evolutionary history of species, helping clarify how

coral families are related or whether scleractinians

evolved before octocorals, for example. In
18

gene has been shown to be the most variable of all octocoral mitochondrial genes sequenced to date

(France and Hoover 2001; Sanchez et al. 2003b). This gene is likely to be useful for phylogenetic

studies of octocorals but has not yet been found in scleractinians.

Other genes that are being tested for applicability in phylogenetics and population genetics for both

octocorals and scleractinians include the mitochondrial DNA control region, internal transcribed spacers

(ITS), and several introns (see Van Oppen et al. 2000 and reviewed in Shearer et al. 2002). Of these,

ITS is the most variable, but can be difficult to align. It has been widely used in studies of shallow-water

scleractinians (Medina et al. 1999; Fukami et al. 2004; Vollmer and Palumbi 2004) and has been used

in one study of hybridization in shallow octocorals (McFadden and Hutchinson 2004). However,

Sanchez and colleagues (2003b) found that ITS sequences were not alignable across families of

octocorals, and they suggest that this gene may not be useful for octocoral phylogenetics.

Population Genetics

Population-genetic studies are often used to infer dispersal patterns in organisms for which movement of

larvae or adults cannot be tracked or inferred. These methods should therefore be particularly useful in

understanding the ecology of deep-sea organisms but have rarely been applied in deep-sea settings.

The low variability in mitochondrial genes that plagues coral systematists is even more of a trial for

population geneticists. Mitochondrial genes such as COI and 16S rRNA are often used in populationgenetic studies of other invertebrates but may be invariable at the genus level in corals. Early populationgenetic studies of shallow-water corals used allozyme markers (Abbiati et al. 1993; Burnett et al. 1995;

Hellberg 1994, 1995; Benzie et al. 1995; Ayreand Duffy 1994; McFadden 1997). Allozymes are allelic

forms of enzymes that can be separated using electrophoresis. They are inexpensive and can be

informative for population-genetic studies but do not have the same resolution as the other methods

discussed here. DNA fingerprinting of individual organisms using amplified fragment length

polymorphisms (AFLPs) has also proven useful in coral population genetics (Barki et al. 2000);

however, this method has not been widely applied.

Microsatellites are the most promising markers for coral population-genetic studies. Microsatellites are

segments of DNA that consist of repeated units of short (di- or tri-nucleotide) sequences.

Microsatellites are highly variable and are codominant, meaning that both alleles at a locus can be

scored, facilitating the estimation of population-genetic parameters (Roderick 1996; Palumbi 1996;

Beaumont and Bruford 1999; Chakraborty and Kimmel 1999; Barki et al. 2000). Microsatellites have

been shown to be sufficiently variable in corals to be useful for population-genetic studies, and they are

currently in use in many studies of both shallow and deep-sea corals (Maier et al. 2001; GutierrezRodriguez and Lasker 2004; LeGoff-Vitry et al. 2004b; Magalon et al. 2004; Shearer and Coffroth

2004; Baco and Shank 2005; Baco et al. in prep). Though useful, microsatellite loci must be acquired

for each species under study, and the loci are rarely usable across taxa, even within a genus. In addition,

microsatellites require considerable startup time and funds to develop.

Sampling Requirements for Genetic Analyses

Phylogenetics: 1-3 individuals of each species of interest, plus an outgroup species.

Population Genetics: 30 or more individuals of the species of interest from as many different sites or

populations as possible.
19

Handling procedures adopted by the Mountains-in-the-Sea Research Group

by Les Watling

Fig. 18a and 18b. The Mountains-in-the-Sea Research

Group in action

The Mountains-in-the-Sea Research group

got its start in the canyons of

the northeastern United States, where we were first confronted with octocorals of various kinds, but

most of which were pretty hardy. During the course of our subsequent expeditions to the New England

seamounts of the North Atlantic, we have come to recognize that certain of the octocoral families have

to be handled differently from others. Handling issues are especially important when bringing specimens

from the cold deep sea through warm temperate or tropical surface waters. Deepwater temperatures

can drop to 2 oC. However, the collection and handling procedures outlined below can be modified

somewhat when dealing with surface waters that are approximately the same as those at the collection

depth. We do not yet know much about the temperature tolerance of octocorals and antipatharians

living below normal scuba diving depth, so our guiding principle has been use methods that cause as little

shock to the organism’s tissues as possible.

Collection Procedures

Manipulator arm. We found that if octocorals and black corals (and perhaps other delicate specimens

as well) are the only items to be collected with the manipulator arm of the submersible or ROV, the

standard three-fingered claw should be replaced by a cutting and gripping style claw. The one designed

for us by the Institute for Exploration can be seen on NOAA’s Ocean Explorer web site. It consists of

two stainless steel angle-iron pieces with two Nalgene tubing pieces arranged so that as the sharpened

edges of the opposing angle-irons cut the coral stem the tubing acted to grip the cut piece.

Collection boxes. When there is a possibility that the specimens will be recovered through surface

waters that are more than a few degrees warmer than the temperatures experienced at the collection

site, we try to use tightly sealing insulated bio-boxes. In some cases we have not been able to have all

insulated boxes, so we have experimented with which corals put in the non-insulated boxes
20

 and still be useable when recovered. Bamboo corals are the least likely to be useful for any

morphological or genetic work if they spend much time at temperatures much above those where they

were collected. Note here also that recovery times for the submersible or ROV can be quite long. For

example, after Alvin enters the warm surface waters, another 45 minutes can pass before the sub is

recovered and safely secured and you have access to the specimen boxes. With ROVs, this time can be

shortened somewhat, but can still seem excruciatingly long.

Oxygenation. Some ROV dives are as long as 17 hours or more. In that case the lid of the bio-box

should be opened every 2 hours or so in order to oxygenate the water in the box. Remember that some

deep-sea waters are low in oxygen as it is, so specimen deterioration can occur in the box. In a few

cases we have seen that associates of the corals will abandon the host when conditions get bad in the

box, but for the most part the associates hang on to the end.

Mucus production. Many octocorals secrete copious amounts of mucus when stressed. As a result we

try to be careful about what families of octocorals and black corals go in a bio-box together. For

example, antipatharians go into a box of their own whenever possible. Bamboo corals, large primnoids

(such as Primnoa resedaeformis), and the very large Paragorgia spp. are pretty good mucus

producers. Small primnoids, chrysogorgiids, plexaurids, and other small octocoral species do fine

together in one box.

Video data collection. Before collection we take very close video of the specimen (using lasers for

scale), especially of the colony base which is often not recovered. Longer term close-up video also can

be used to show the tentacle pinnules (which can often be counted if the light is right), and to see how

any associates living on the coral might be behaving. In practice, we have not been able to get more than

15 minutes of close-up video, however, most likely because the photons from the lights begin to heat the

polyps and they start to close or show other signs of being stressed. Video of the surrounding habitat,

including small-scale geological features, surficial substrate type, macro-associates and other sedentary

fauna should also be recorded, as described in the following section

Record keeping. Every place where the ROV or submersible stops is given a station number and the

specimen(s) collected are assigned individual specimen numbers. For example, in our system, KEL

408-1, is interpreted to mean Kelvin seamount, 4th dive, 8th station, 1st specimen. With this system we

can determine exactly which video we need to view to see the collection of the specimen. We also note

the bio-box into which the specimen was put and make a “map” of each bio-box with tentative

specimen names, small drawings of the specimen showing its shape, and the order in which the

specimens were put into the box. These details will be very important in sorting out the 15-20

specimens in the boxes when everything gets to the surface.

Shipboard Procedures and Other Considerations

Immediate storage. While the ROV or submersible is in the water, a sufficient number of 5-gallon

buckets of seawater are chilled to the at-depth temperature. Each bucket is labeled for a single

specimen. As soon as the specimens can be removed from the bio-boxes, they are put into the buckets

and taken for storage in the cold room.
21

Specimen deterioration. We have seen that specimens in the different coral families deteriorate at very

different rates. Bamboo corals often break down very quickly and are handled first, followed by

plexaurids such as Paramuricea sp. and its close relatives. Chrysogorgiids last for a moderately long

time (although many polyps die extended and often with tentacles extended as well) as do Paragorgia

and its relatives. Deterioration of bamboo corals can be seen in the tendency for the polyps to begin to

slough off the axial skeleton. In Paramuricea and relatives, the tentacles begin to turn black within

about 20 minutes of being put into the buckets.

Fixation. There has been a lot of discussion on various lists about using formalin, especially with

respect to deterioration of sclerites. While there are many drawbacks and health issues associated with

this fixative, there are many benefits, the most important of which is the ease with which delicate tissues

can be rapidly fixed. Fortunately bamboo corals have easily-penetratable tissues, which may also be

why they deteriorate so rapidly, even when handled with the best of procedures. As a result we have

adopted the following quick fixation procedure to be used for all octocorals with especially delicate

tissues, such as the bamboo corals and chrysogorgiids. One or two buckets of 4% formalin-seawater

are prepared and stored capped in a fume hood. After the specimen has been tagged with its unique

identifier, it is put into the formalin bucket for about 12 hours. Following the formalin immersion it is

rinsed in another bucket containing seawater and then transferred to a bucket or bag of 70% ethanol.

The result is specimens that will be useful for taxonomic studies for a long time. We’ve seen that these

formalin fixed specimens show little contraction of the calyx and the relationship of the sclerites to the

coenenchyme is preserved, at least macroscopically.

Other data collection. Fixation issues aside, once the specimens are safely stored in the cold room,

each specimen goes through the following procedure before being finally stored for later use. The order

in which the first three of these items actually occur depends largely on the family of coral being dealt

with, the length of time since collection, and etc.

1. Labeling. Each specimen is labeled by writing its identifier, collection date, and tentative

name, onto a water-proof heavy paper which is tied to the specimen with monofilament line.

2. Genetic sampling. Before the specimen is handled by too many pairs of hands (all equipped

with suitable gloves), pieces are set aside for genetic analysis. Usually we try to have a small

ice-filled tray in the lab into which can be set plastic dishes (such as sandwich containers

available from any supermarket) containing cold seawater. The genetic samples can be stored in

these small dishes until they can be dealt with.

3. Photography. The entire specimen with its label and a ruler for scale is photographed,

usually on a black cloth, but we have also used a gridded board. Close-up photos of special

features or associates are also taken. We use a 5-megapixel camera with close-up capability.

Several pictures are taken to be sure that at least one could be used for publication of the

morphological description of the species.
22

4. Reproduction. Sections of the coral for reproductive analysis are then removed. Details of

the methods for fixing and preserving tissue for reproductive analysis will be dealt with

elsewhere, or interested persons can contact me.

5. Associates. All associates are then removed, if possible, and fixed appropriately. The

container(s) in which these animals are stored is given the same identifier as the coral from which

they were taken. Associates are not removed where excessive breakage of the coral specimen

will result. Fixation using formalin (4%) or alcohol follows normal techniques for the various

phyla. For example, polychaetes should be fixed for 24 hours in formalin, then rinsed and stored

in 70% ethanol, whereas ophiuroids, if they are to be used for taxonomy, should never be

formalin fixed. I suspect, however, that as with the octocorals, very short fixation with a weak

formalin solution will probably not ruin the carbonate structures needed for taxonomic analysis.

We’ve used other fixation protocols, depending on specific uses of the organisms.

Final storage. After all of the various bits and pieces of the specimen have been dealt with and the

specimen has been properly fixed, it is prepared for storage aboard ship and made ready for shipping.

Specimens are handled according to their special characteristics. Small fan-shaped species are stored in

ethanol in heavy-duty zip-loc bags. A group of 10 or so of these bags is then put into tightly sealing

chemical storage boxes purchased from an industrial supplier. If the bags develop micro leaks, which is

frequent, especially with small primnoids and many of the bamboo corals, the box will contain the

leaked alcohol. Some specimens are double-bagged. Larger specimens are loaded into larger bags and

stored flat in larger boxes, but it is not possible to get really large (about 44 inches long) boxes with

tightly sealing lids. In these cases we have often soaked the specimen in alcohol in a box or bucket for a

day or so, then put the specimen in a bag with a small amount of alcohol and removed as much air as

possible. (As part of another project in Alaska a vacuum system was used but we haven’t evaluated the

results yet. Gradually reducing the amount of alcohol facilitates shipping as only small quantities are now

allowed.) Very large specimens are stored in their own buckets with sealable lids. These buckets are

maintained full of alcohol until shipment.

Figs. 19a and 19b. Roundnose fish and Paragorgia sp. on a New England seamount in the eastern North Atlantic.
23

Documenting deep-sea coral collections

In situ video and photography is often available to modern deep-sea explorations. Most submersibles

and ROVs are equipped with 3 chip digital video cameras and high resolution still cameras. Video and

photography are particularly useful for sharing information between scientists, for tentative species

identifications, and for public outreach efforts. Video documentation has the additional benefit of

recording community structure, e.g. target species density along a transect, and associated commensal

species. Still photo documentation is most useful for tentative species identifications and for publications.

High quality images can depend on several things. The most important are proper exposure, and proper

resolution. Too much light (often from a flash) can result in over-exposure. Too little light will result in

underexposure, and a grainy image with poor detail. Angle the lights 45 degrees downward to avoid

backscatter from plankton. Try and use your available light judiciously, and get as close as possible to

your subject. Still image captures from video are limited to the scan resolution of the video and the

software. Some video captures are very low resolution, e.g. 72 dpi. This is suitable for internet

distribution. Print quality (300 dpi) in situ images may require a still camera, or a “still photo” setting on

the DV Camera.

Sseparate video and photo protocols are provided. The video section will discuss camera, lights, and

media format. The video section will also present some proven video transect methods and some

recommended camera angles. The photo section will discuss “life cycle” photo documentation so that

your in-situ photo stills can be matched to dried specimens on deck, and to live polyps under the

microscope.

Video protocol

1. Camera Quality:

a. 3 chip (CCD) video cameras are superior to one chip video cameras.

b. Low light (SIM) cameras are grainy, and poor. HDTV cameras are the best available.

2. Lights:

a. Quality: All lights have a different color temperature, and the camera setting must match that

temperature using the white balance. Daylight is 5200 K, tungsten light is 3600 K.. Xenon lights

are approximately 4200 K.

b. Quantity: 1000 watts of Xenon light are best

c. Position: Lights positioned behind armatures cause shadow, and/or underexposure because they

reflect light into the camera iris. Position lights high and to the side, angled downwards.

3. Archival Media Quality:

a. HD, DVCam, Digibeta, Beta SP, and Mini-DV are archival quality video tape formats.

b. These are magnetic tape mediums, and will degrade within 10-15 years.1

4. Archival digital resolution:

a. Broadcast quality digital video (.mov, .mpg, .mp4) is 720 x 480 pixels

5. Distribution digital resolution:

a. Medium: 320 x 240 Compressed video is most suitable for internet distribution

b. Small: 160 x 120 is commonly referred to as “postage stamp size”, very small

.

1

 Temperature and humidity are the most important factors to maintaining footage for extended time spans. If you plan

to start a collection, or have an aging one, consider dubbing tapes five to seven years old to a contemporary tape

medium, such as Beta SP, then to a digital medium, such as DVD. If you have 3⁄4” Sony tapes older than ten years you

may wish to consult a restoration facility, such as SPECS Brothers in Ridgefield Park, NJ. www.specsbros.com
24

Video protocol (cont.)

6. Data Burn in

a. Ideally, data will be burned into one camera only,

including dive number, depth, and geoposition. A

second camera should have clean, high resolution

images for broadcast and distribution.

b. In many cases a second camera may not be

available. Some recorders can turn the overlay

information ‘off or on’. Harbor Branch

Oceanographic Institute (HBOI) uses the overlay

‘on’ initially to document the overall habitat while

they verbally describe a sample, and then turn the

overlay ‘off’ for highest quality video of the

specimen.

7. Camera angles and recommendations

a. Wide, medium, tight

i. Record a wide view of the community

1. How dense is the community?

2. Is it monotypic or diverse?

3. Are there conspicuous fish

 or invertebrates nearby?

ii. Record a medium view of the colony from

base to tip

1. To what substrate is the base

attached?

2. Do you notice any cryptic species

or epibionts?

iii.Record a close-up view of polyps,

 branching morphology, and commensal

 species

b. Reasons for getting close

i.Pro - better light

ii.Pro - less apparent movement in a wide lens

iii.Pro - better view of polyps and

 commensals

iv. Con - potential for turbidity

v. Con - potential for damage to colony

c. Reasons for keeping your distance

i. Pro - Minimally invasive

ii. Pro - Greater range of possibilities for

wide, medium, tight shots.

iii.Con - Less light, grainier image.

iv. Con – More apparent camera movement

when “zoomed in”

Fig. 20a. Wide community shot, 6 ft. sponge!

Fig. 20b. Wide colony shot with burn in.

Fig. 20c. Medium shot. Note associated species.

Fig. 20d. Close up with associated species and

10 cm redlaser reference for scale.
25

Video protocol (cont.)

Video transect types3

:

Line (Strip) transectVertical line transects (Fig 21a) are best for

rapid biological assessments of large features,

such as seamounts. These start at the base of

the feature, and almost guarantee dramatic

changes in environmental conditions.

Vertical line transects can be repeated around

a feature. They also help discern vertical

zonation patterns, and the results are readily

quantifiable.

Grid TransectGrid transects (Fig 21b) can be very time

consuming, but they can also cover a large

area systematically. The results are readily

quantifiable. The length of the transects, and

the offset are usually predetermined.

Grid transects may be oriented parallel to

particular isobaths, or perpendicular to those

isobaths. This technique is very powerful

when combined with an opportunistic

sampling methodology to verify video

identifications.

Radial transectRadial transects (Fig 21c) start from an

arbitrary or randomly selected

center. These are best for smaller features,

or low relief mounds. A radial transect can

help determine the limits of a particular

community when the size of that community

is unknown. All axes need not be the same

length.

3

The analysis techniques described here are modified from those developed for the

Great Barrier Reef in Australia (after Osborne, K., and W. G. Oxley. 1997.)

Fig. 21a-c. Video transect types
26

Still image protocol

Digital still cameras provide much better resolution than video still frames. These two mediums are not

interchangeable. Print quality images are 300 dpi. Video still frame resolution is derived from the number

of horizontal lines in the video image. On a 3-chip camera, this amounts to about 72 dpi. This is good

enough for internet distribution, but not for print. Newer High Definition (HD) video cameras can

produce frame grabs at high resolutions more suitable for print. Unfortunately, these cameras are not yet

widespread. Plan to take your best pictures with a digital still camera, rather than depending on video

‘frame grabs’.

The most useful method of

documenting biological collections is

called “living specimen photography”.

 First, document the specimen in-situ,

(Fig. 20) with wide shots, medium

shots, and close-ups of polyps,

before you collect it. Once you bring

the colony on-deck, place it in

refrigerated seawater and cut a small

branch from the colony to examine

and photograph the extended polyps

through the oculus of a dissection

microscope. Then photograph the

colony.Sample numbers and scales

can be useful additions to the image.

Fig. 22. Scale is difficult without clues, like laser points, or the crabs

shown here. This primnoid colony is ~ 4 feet tall.

Small extended lenses of modern digital cameras perform nicely with a microscope. This procedure

takes time. Polyps can remain retracted for hours, shocked by temperature and lighting differences.

Some polyps may appear in a bucket of seawater as it sits in a dark cooler for a while. To complete the

photo cycle, photograph the entire colony again with a scale reference, so that you can compare the

appearance in situ to the appearance ondeck. You may wish to pack some

duvatine, a light absorbing black fabric,

and some 2’ x 3’ dark colored felt material

to use as a plain background. Some

colored thin foam mats are water repellant.

Fig. 23. The dried skeleton of Keratoisis

flexibilis photographs nicely on a neutral grey

background. Scale is again absent, but the image

is perhaps better for it. Image ~ 25cm.
27

John Reed's August 22 article on Stetson's Reef from Islands in the Stream 2002 expedition has some

terrific images, as does the entire 2003 Life on the Edge Expedition, courtesy of Art Howard from

NAPRO Communications. Notice that these examples have plain, appealing backgrounds and sharp

focus (Fig. 22). A tripod and good lighting will improve your depth of field, your camera stillness

(unless your boat is rocking!) and your focus. Small flat plastic rulers are important for scale. Coins

are also useful (see Fig. 9). Simple labels, like a dive number or sample number, should be included.

Place these labels so they can be easily cropped out of the image if necessary. You can study these

examples of some high quality digital photography at NOAA’s Office of Ocean Exploration website:

http://oceanexplorer.noaa.gov/

1. Image Quality

a. Printed images or CDROMs of still images may accompany a sample

b. Print Quality Digital Files: 300 - 600 dpi .tiff, .png, or .jpg image files

c. Distribution Quality: 72 dpi .jpg image files

d. Low resolution still images may be collected ex post facto from video

2. Camera Framing and zoom control (see insets on page 18)

a. Community Image- the widest shot, includes organism, substrate and associated species

b. Wide Shot- frame the entire colony as best you can. (Fig. 21)

c. Medium Shot- branching morphology and associated species

d. Close-Up Shot- epiphytes

e. Reference – Any reference for size and scale, e.g. ROV claw or laser scale

3. Camera angle

a. For transects to document habitat cover (%), you can have the camera angled at 90

degrees to the bottom (plan view) or 30-40 degrees ahead for better estimates of colony

height. Try to keep parallel lasers in view for scale. ALVIN uses two laser pointers set

10cm apart.

4. Data Burn In

a. include date and time, dive number, depth, and geoposition in metadata

b. alternatively, enter this data into the image itself

5. Practice Living Specimen Photography

Fig. 24. Alaska’s Aleutian Islands have diverse

and colorful coral gardens deeper than 300 m.

Commercial bottom trawl fisheries can damage

these gardens
28

Fig. 25b. In-situ medium shot. Polyps extended.

Fig. 25d. On-deck medium shot. Polyps retracted

Fig. 25f. Dissecting scope close shot (40x)

Fig. 25c. On-deck wide shot on a measured grid

Fig. 25e. Dissecting scope medium shot (10x)

Fig. 25. Example of the sequence of photographs documenting a deep-sea coral. Photographs are taken from the

deep sea (a,b), on the deck of the boat (c,d) and under a microscope (e,f).1

Living specimen photography

Living specimen photography helps increase our national capacity to document deep-sea coral diversity.

The resulting images can be used by subsequent expeditions to identify coral colonies in-situ, reducing the

need to collect new specimens, reducing time and effort for future expeditions, and facilitating broader

scale habitat characterization efforts. The method photo-documents the specimen in-situ with a wide

shot, a medium shot, and a close up of the polyps, collects a branch or colony, then reproduces these shots

on deck with a scale, and photographs polyps in the wet lab with a microscope. This is a time consuming

process, but the results are useful. Pictures show Swiftia sp. in the Gulf of Mexico.

Fig. 25a. In-situ wide shot. Nearby colonies evident.
29

Collecting

There is a broad range of opinion about how much coral to collect, depending upon the intended

purpose of the sample. Most museums would prefer an entire colony, especially if the specimen is a new

species, or intended for display. These can be large, though, and difficult to ship. For age and growth

studies, the holdfast of a large colony is most important. For species identifications based on gross

morphology, a small branched segment will often suffice, as described in the chapters by Sanchez and

Cairns. For molecular studies, a small amount of preserved tissue is most useful, particularly if tissue can

be collected from several colonies.

One thing deep-coral researchers can agree upon is that photographs and videos alone are not

particularly useful for species level identifications. Photographs can help, of course, especially when

accompanied by a physical specimen, and referenced with a biogeographic database. Photographs are

often suitable for genus and family level identifications.

There is also a broad range of opinion about how to preserve a sample, and this is discussed on

following pages. For some corals, like bubblegum and primnoid corals, dry samples can be useful.

Dried skeletal materials of ahermatypic scleractinia, antipatharians, and bamboo corals are also useful

for radiocarbon analyses. Some biologists feel that dry samples should be made only when absolutely

necessary; for example, when you don’t have room to store the whole specimen. Wet samples collected

by the Mountains in the Sea Research Group are not small, on purpose, because they don’t want to

lose information that only wet samples can provide. Large wet samples will benefit the most future

studies, but 6-10" branched segments are usually sufficient for species identification.

Your study organism may have the opportunity for regrowth if you leave the organism in place, and take

only a branch. This is particularly relevant to cases where repeated samples of a single species are being

collected. Selective clippings are the most responsible way to perform good science without impacting

a localized population. Try to take only a clipping from the largest specimen. The largest colonies may

be the most productive, and should be left alive in situ. It is preferable to collect your sample from a

dense aggregation. Solitary colonies may be pioneers that facilitate more settlement, and therefore more

future habitat.

Permits

Orders Scleractinia and Antipatharia, and Family Stylasteridae are listed on CITES Appendix

II. Collection from U.S. waters and subsequent transport within the U.S. does not require a

CITES permit, transport of these corals across international borders requires an export permit

from the CITES Management Authority in the country of origin. Collections in state waters (up

to 3 miles) need a state permit, those in Federal waters need a Letter of Acknowledgement of

scientific research, which is available from regional NMFS offices. To collect from Marine

Reserves, you need special permission from the reserve management authority. These permits

are granted by regional fisheries science centers. A list of these centers is available here:

http://www.noaa.gov/fisheries.html.
30

Preserving

When brought aboard, octocorals can exhibit a foul odor. Be prepared. Always work with a ventilation

hood. Field systematics can be perfomred with a careful review of the polyp and sclerite morphology

under a microscope, in reference to morphological and photographic keys. Further laboratory analyses

(e.g. SEM microscopy and molecular classification) will require well preserved tissue samples. The

preservation medium depends on your research intentions. These vary in toxicity from 95% Ethanol

(grain alcohol, a preservative) to Formalin (a highly toxic, carcinogenic, all purpose fixative). Tissue

fixation is not absolutely necessary for species identification. Preservation in 70- 95% ethanol is usually

sufficient. To fix specimens (i.e., change their structural properties) for molecular work, histology, or

long-term preservation you need a fixative.

These protocols for preservation have been established largely with Gorgonacea, branched octocorals,

in mind. The techniques outlined in the section on handling procedures by Watling all work on both

black corals and zoanthids. The preservation method you choose depends upon your purposes, most of

which are outlined here.

Dried Museum Specimen

a. Before drying a specimen, preserve reference tissue from a mid-size branched segment.

b. Branches range from delicate/fleshy (black coral) to thick/spongy (Paragorgia sp.) to bony/

oozing (bamboo coral). Some corals clean up nicely, e.g. bamboo corals.

c. Hang the specimen to dry—outdoors, if possible — in a splash free environment. Avoid laying

specimens on deck due to safety concerns. If you must lay specimens on deck, secure the area,

obtain permission, and take appropriate precautions.

d. If you must dry a specimen, especially because of size, you may soak the specimen in formalin

over night, then stand upright in a fume hood for two days before wrapping in plastic for

storage. Putting the whole specimen in a cardboard sleeve (made from boxes) will protect

against breakage.

Tissue Fixation

a. A general comment on fixing tissue for analysis, these pieces should be small enough to allow full

penetration of the preservation medium - no more than 2cm thick. As with any fixative, use

approximately a 1:10 ratio of tissue to fixative.

b. Formalin is readily available and cheap and can be used at 5-10 percent. Full strength formalin

is a 37 percent solution of formaldehyde. To make a 10 percent solution, dilute the 37 percent

formaldehyde (formalin) 1:10 (not 1:3.7).

c. To reduce calcium dissolution, formalin can be buffered by using seawater as the dilutent and/or

addition of borax crystals. Samples can be held indefinitely in formalin, but calcified samples

(even if buffered formalin is used) should be rinsed briefly and post-fixed in 70% ETOH after

24 hours.

d. You can ship small quantities of ethanol (<30ml) without special paperwork. Some samples can

be wrapped in an ethanol-soaked paper towel in an airtight container. On reaching the lab,

specimens can again soak in 70% ETOH or back into 10% formalin, depending on their

ultimate purpose. The use of neutralizing solutions, which are commercially available from

scientific catalogues, can solve waste disposal problems.
31

Tissue preservation

a. Do not let specimens dry. Keep them cool in native seawater.

b. We recommend 70-95% ethanol or analytical absolute alcohol to preserve tissue, not denatured

alcohol (which is about 100% but it has some nasty stabilizers).

c. Formalin is not universally recommended for preserving octocorals. Formalin is toxic,

carcinogenic, difficult to transport, and degrades DNA. As above, formalin can be used briefly

as a fixative to get good tissue preservation without degrading sclerites.

d. The smallest samples should be in 30 ml vials (e.g. blue-tip vials from Fisher). For molecular

work, use 12ml and 3ml plastic vials with color coded caps.

SEM Analysis

a. In the particular case of octocorals, samples for SEM analysis are usually digested in bleach

(e.g., Clorox) and then coasted with gold. You should preserve material in ethanol or let it dry. It

is easiest to dry the material, but preserved tissue will provide DNA.

b. Glutaraldehyde is also used as a fixative for SEM preps, and tissues are transferred to buffer

after an hour in fixation. Glutaraldehyde is also very toxic material.

DNA Analysis

a. DNA samples need to be fixed as soon as possible after collecting. Do not let specimens dry

before wet fixation.

b. Do not use your hands to collect branches for DNA preservation. Even if you wear gloves, use

forceps and a scalpel to collect the branches for DNA preservation. Using a clean kimwipe, blot

excess water from the tissue just before adding it to preservative. Clean (with ethanol) or

change the forceps and scalpel after every collection.

c. Preservation Options for DNA Analysis:

1. Frozen samples

i. Samples frozen in liquid nitrogen work best, as long as they can be kept frozen

ii. An ethanol/dry ice bath should also be sufficient

iii. Store at -20 to -80 degrees Celsius

iv. Secure against power failures and minimize freeze/thaw episodes in the lab.

2. Ethanol (95 percent)

i. Ethanol will suit both tissue and DNA preservation but may not be ideal for

either.

ii. Use of ethanol reduces the number of solutions brought on board.

iii. DNA degrades in ethanol-preserved samples after 5-10 years.

iv. An ethanol concentration of 70 percent may be sufficient but is not

recommended. We recommend 95 percent ethanol if available.

v. In an emergency, ethanol is available as Everclearin liquor stores. Some

twentieth century Caribbean marine biologists used rum for their choice of

preservative!
32

3. Dimethyl sulfoxide (DMSO; 95%)

In a population genetic study with a gorgonian in the Caribbean, DMSO worked 90%

of the time versus 50% of the time using ethanol. DMSO is non-flammable and nontoxic but it penetrates the skin, so take precautions against contact with contaminants or

toxics that may use the DMSO medium to penetrate your skin. DMSO is often used in

medicines as a way to slowly release an active component (sticky patches for sea

sickness, bladder infection treatments). This saturated saline solution is preferred for

fixing DNA, but this is a difficult concoction.

i. DMSO: 250 ml 0.5M EDTA pH=8.0 100 ml DMSO (final concentration 20%)

ii. 105 g NaCl ddH20 for about 400 - 450 ml, Stir a couple of hours, add 10 g of

NACL and stir overnight

iii. Next day complete volume to 500 ml and let the excess salt to precipitate.

iv. 1.5 ml for 0.5 cm3 of sample, use gloves and prevent contact with skin

There is some question about our ability to get good DNA out of samples that have

been stored longer than 5 years in DMSO. Some researchers at the prefer DMDM

Hydrantonin as a non-toxic, non-flammable, long-term solution for tissue preservation.

RNA Analysis

a. RNA is much less stable than DNA. Various RNA specialty companies market buffers.

b. Alhough immediate freezing (but not DMSO) can adequately protect RNA, many researchers

use “RNAlater” (Ambion, Austin Texas, 512-651-0200) to store tissue samples for subsequent

lab work.

c. RNA will degrade after < 1yr, even if stored at –80. if you want to use RNA down the line,

RNA Later is essential.

.

Fig. 26. Primnoid corals in the genus Narella, from Hawaii's deep coral beds.
33

Some preservation techniques for (deep water) coral samples for subsequent molecular

studies: a special supplement from Harbor Branch Oceanographic Institute

by J.V. Lopez

Tissue Storage for Molecular Experiments: To store field collected animals for molecular biology work,

it is important to store the sample such that the DNA is protected from endonucleases, which can

quickly degrade the DNA once harvesting occurs. Gloves further protect samples from degradation.

One option is to immediately plunge the sample into a deep freeze immediately, and in the field

this entails liquid nitrogen (LN2) or an ethanol/dry ice bath. Samples then can be transported in LN2

dry carriers, which though convenient are expensive and must be charged after 1-2 weeks with more

LN2. At destination, most samples can be stored indefinitely in a freezer (-20 to -80oC) for subsequent

DNA analyses. Alternatively, place the sample in DMSO (dimethyl sulfoxide) storage buffer, or place

the sample on a dessicant. Frost-free freezers stay frost-free by going through freeze-thaw cycles, which

is bad for DNA and RNA samples.

DMSO storage buffer (Adapted from Seutin and White, 1991)

20% DMSO

250 mM EDTA

dH2

O (to final volume)

DMSO buffer is SATURATED with NaCl. In practice this means about 210g or more per liter. It takes a

while to go into solution. Make up a 0.5M EDTA (pH 8.0) solution first, then add DMSO and NaCl to half

of this, adjusting the pH to 8.0 as you go - it will help dissolution. Final pH should be 8.0. Place the sample

in a wide mouthed tube (with a lid that seals water tight, such as a 15 or 50 ml conical polypropylene

centrifuge tube; this depends on the size of your tissue specimen), and add enough DMSO storage buffer

to cover the sample. Samples can be transported in this solution and stored at room temp for up to 2

weeks, then should be placed in a 4o

C refrigerator. These samples can then be stored for several months

in the fridge. Dawson et al (1998) gives an excellent evaluation of various other storage techniques, but

supports the DMSO solution as the best for DNA studies.

For subsequent RNA studies, using the frozen tissue protocol will often suffice, or tissue samples can be

placed in “RNALater”, specifically designed for RNA preservation when used with the manufacturer’s

directions (Ambion Inc -www.ambion.com; Austin Texas).

For more details see Baker et al. 1997, France and Kocher 1996, and Lopez et al. 2002.
34

Standard Deep-sea Coral Preservation Protocols Tear Sheet

1. Care should be taken not to contaminate samples. Use rubber gloves and

change them periodically. Use non-serrated forceps to avoid tissue build

up in serrations. Clean forceps between samples.

2. Maintain the specimen in native seawater as long as possible.

3. Transport the specimen to cold storage for handling (if available).

4. Use clean scissors to clip a branched segment with extended polyps into a

petri dish of the coral’s native seawater.

5. Light and photograph extended polyps under the dissecting scope

6. Fix the remaining colony in refrigerated 4% buffered formalin seawater for

~12 hours.

7. Prepare duplicates of each sample, i.e. two vials for each coral collected,

one for a curator and one for research.

8. The size of the jar or vial and the choice of preservative depends on your

needs and your preferences

9. Prepare 50 ml vials with 70-95% ethyl alcohol or 33-50% isopropyl alcohol

solutions made with fresh water, not seawater

10. Prepare 15 ml polypropylene centrifuge tubes with 95% DMSO

11. Place pencil marked label of archival paper inside each vial

12. Use clean scissors to clip one 4 -5 cm branched segment and another

piece of tissue 1-3cm

13. Place 4-5 cm branched segment in each 50ml vial

14. Place 1-3 cm of tissue in each 15ml vial for genetics

15. Change alcohol after two days.

16. Additional material may be dried or frozen.

17. Dried or frozen samples should be bagged in plastic with internal and

external label, and then stapled, taped, or zip-locked shut.

*Modify accordingly. These are only recommendations.
35

Labeling

The significance of proper labels cannot be underestimated. If you are sending your samples to a

scientific institution or laboratory, all the hard work of collection may be lost to dissolved ink or illegible

handwriting. Furthermore, improper labels can be frustrating to those trying to help you. Researchers at

the Smithsonian Institution note that externally labeled vials tend to lose their markings. The best labels

are on paper inside the vial, in the preservative, clearly written in pencil. You may wish to keep

a sample log, and reference each entry to the label in the vial. Wet samples should be related to dry

samples. Vials should also be labeled on the outside on the cap and the body. Use only solvent

resistant markers for the exterior label. These vials should be wrapped in parafilm before shipping.

Label Information

1. Use pencil on a small strip of paper approximately 1⁄2” by 4"

2. Paper should be either 100% rag bond, Nalgene (TM) or Resistal (TM) contact: University

Products 517 Main Street, P.O. Box 101, Holyoke, MA 01041-0101.

3. Write the date, cruise name, ship, researcher, station number, and sample number in clear

block letters include geoposition (longitude and latitude), place name, depth in meters, related

wet or dry samples, collection method, and substrate (Fig. 24 and 25).

4. Geoposition should be

a. Degree/Minutes/Seconds (DMS) e.g. 32 15’ 43"N, 80 25’ 18"W or

b. Degree/ decimal minutes (DDM) eg 32 15.23 N

c. Decimal Degrees (DD) to three decimal places e.g. 32.262 N, -80.421 E

d. 1 degree = 111km, so 2 decimal places is only accurate to within 1km

e. To convert DMS to DD, DD = D + M/60 + S/3600

f. To convert DDM to DD, DD = D + DM/60

5. Depth should be clearly defined in meters (vs fathoms, feet, km, nmi)

6. Depth may be a range of values due to gear type

Fig 27. This label shows collection information in pencil, not

ink. The paper is durable and archival. Rubber gloves would

help avoid DNA/ RNA contamination.

Fig. 28. 50ml vials hold small branches in

ethanol, but legally require extra paperwork

to ship overnight. You may drain these to

ship them, and refill them later at the lab.
36

Shipping

Packages shipping biological samples preserved in formalin and ethanol are subject to stringent rules and

regulations. You must be certified to ship hazardous materials. It is illegal to ship hazardous materials

without proper certification. The techniques and procedures described below are excerpted from a

49CFR/ICAO/IATA Hazardous Materials Transportation Certification class.

To summarize, preserved samples must to be triple-packed in specific container types with limited

quantities of fluid in order to meet the “Dangerous Goods in Excepted Quantities” provision of

International Air Traffic Association (IATA) and US Department of Transportation (DOT). This protects

the safety of the package and the individuals handling it.

Tissue samples preserved in ethanol must be packed in a sealed primary package (like a vial or jar) with

<30ml of fluid. These must be contained in a sealed secondary package (like a plastic bag) with <500ml

total fluid. A bag with 10 50ml vials (50x 100 = 500ml) will qualify as "Dangerous Goods in Excepted

Quantities" when it is packed and sealed in sturdy cardboard box. The box must be no smaller than

100mm by 100mm on two sides. You can reduce the volume of ethanol in the primary package by

wrapping a sample in cheesecloth or paper towel, allowing this to draw the preservative.

“Dangerous Goods in Excepted Quantities” packages must be labeled on the waybill and on the box. A

“Dangerous Goods” label must be filled out and attached to the side. Check the box indicating Ethanol is

Class 3 material. The UN number is 1170. The “Dangerous Goods” labels found at the websites below

can be used if printed in color, no smaller than 100mm x 100mm.

If you need to ship large specimens or large numbers of specimens, be sure to check with the person at

your institution responsible for shipping dangerous goods. One can ship ethanol-preserved material

under the “limited quantities” designation if the inner containers hold 0.5 L or less ethanol and the total

package contents are 1 L or less. There are different labeling requirements, etc., but otherwise the

regulations are not much more onerous than for the “excepted quantities” designation.

These websites describe shipping regulations for biological specimens in great detail.

Natural History Museum of LA County http://collections.nhm.org/library/hazmat:hazmat

This comprehensive and user-friendly website compiled by the NHM Marine Biodiversity Processing

Center addresses invertebrate specimens directly, with links to printable labels for shipping packages.

University of New Hampshire http://www.unh.edu/ehs/shipping/Excepted-Quantities-Guidelines.pdf

This brief .pdf document by Andy Glode at the University of New Hampshire - Office of Environmental

Health and Safety (UNH-OEHS) provides a clear explanation of primary, secondary, and outer packaging.

It also includes a printable Dangerous Goods label of appropriate size (100mm x 100mm).

Stanford University http://www.stanford.edu/dept/EHS/prod/researchlab/bio/Shipping_Guide.pdf

Stanford University’s “Shipment of Biological Materials & Dry Ice Reference” is an extensive and

detailed .pdf document created by trained authorities Andy Glode & David R. Gillum, UNH-OEHS.
37

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44

Image Credits

Alvin group, Wood’s Hole Oceanographic Institution, NOAA, Cover foreground

and the Gulf of Alaska Seamount Expedition Science Party Fig. 2 and 20A-D

Amy Baco-Taylor; Thomas Shirley; pilots T. Kerby and M. Cremer; and NOAA Fig. 26

E. J. Brill publishers, The Netherlands Fig. 11A – 11C

Sandra Brooke , Oregon Institute of Marine Biology Fig. 16A-B

Stephen Cairns, Smithsonian NMNH Fig. 13, 14

Phil Colla, Ocean Light International Cover background

C Robert Delfs, www.deepseaimages.com Fig. 6

Peter Etnoyer, Harte Research Institute Fig. 1, 2, 3, 4, 16C, 23B-23F, 24, 25C-F, 27, 28

Gulf of Mexico Deep Habitats Research Team, NOAA, and Sonsub Innovator Fig. 25A-B

Harbor Branch Oceanographic Institute Fig. 23

Alberto Linder, Duke University Fig. 24

Mountains in the Sea Research Team, Institute for Exploration Crew, Fig. 5, 17, 19A-B

and NOAA

Olympic Coast National Marine Sanctuary and NOAA Back cover

Pal Mortensen, Department of Fisheries and Oceans Fig. 15

Northwestern Hawaiian Islands 2003 Expedition Science Party and NOAA Fig. 22

Diana Payne Fig. 18A

John Reed, Harbor Branch Oceanographic Institute Fig. 12

Juan Armando Sanchez, Universidad de los Andes Fig. 7, 8, 9, 10

Anne Simpson Fig. 16D

Gary Williams, California Academy of Sciences Fig. i

Dave Wright Fig. 18B
45

Appendix 1. Classification of the Deep-sea Corals

A list of 48 families of hexacorals, octocorals, and calcified hydroids with species known to occur

deeper than 200m. Common names are in parentheses.

Phylum Cnidaria (=Coelenterata)

Class Hydrozoa

Subclass Athecatae

Order Filifera

Family Stylasteridae (hydrocorals, lace corals)

Class Anthozoa

Subclass Alcyonaria (=Octocorallia) (octocorals)

Order Alcyonacea

Family Clavulariidae

Family Alcyoniidae

Family Nephtheidae

Order Gorgonacea * (gorgonians)

Family Anthothelidae

Family Paragorgiidae (bubblegum corals)

Family Coralliidae (precious corals)

Family Keroeididae

Family Acanthogorgiidae

Family Plexauridae

Family Gorgoniidae

Family Ellisellidae

Family Dendrobrachiidae

Family Chrysogorgiidae (golden corals)

Family Primnoidae

Family Isididae (bamboo corals)

Order Pennatulacea (sea pens)

 Family Veretillidae

Family Echinoptilidae

Family Kophobelemnidae

Family Anthoptilidae

Family Funiculinidae

Family Protoptilidae

Family Stachyptilidae

Family Scleroptilidae

Family Chunellidae

Family Umbellulidae

Family Halipteridae

Family Virgulariidae

Family Pennatulidae
46

Appendix 1. Classification of the Deep-sea Corals (cont.)

Subclass Zoantharia (=Hexacorallia)

Order Zoanthidea (zoanthids)

Family Parazoanthidae e.g., Gerardia (gold corals)

Order Antipatharia (black corals, wire corals)

Family Antipathidae

Family Cladopathidae

Family Schizopathidae

Family Leiopathidae

Family Myriopathidae

Family Aphanopathidae

Order Scleractinia (stony corals)

Family Caryophylliidae e.g., Lophelia

Family Dendrophylliidae e.g., Enallopsammia

Family Flabellidae e.g., Flabellum

Family Turbinoliidae e.g., Sphenotrochus

Family Rhizangiidae e.g., Astrangia

Family Fungiacyathidae e.g., Fungiacyathus

Family Oculinidae e.g., Madrepora

Family Micrabaciidae e.g., Stephanophyllia

Family Pocilloporidae e.g., Madracis

Family Guyniidae e.g., Guynia

Family Anthemiphylliidae e.g., Anthemiphyllia

Family Gardineriidae e.g., Gardineria

*Not all Taxonomists recognize the Order Gorgonacea, some believe there is a continuous variation from

the Alcyonacea through the Gorgonacea
47

Appendix 2. Deep-sea coral Panel of Experts (DSC-POE) protocol review team

Judith L. Connor

Director of Information and Technology

Dissemination

Monterey Bay Aquarium Research Institute

7700 Sandholdt Road

Moss Landing, CA 95039 USA

Phone: 831- 775-1728

conn@mbari.org

Peter J. Etnoyer

Research Associate

Harte Research Institute

Texas A&M University-Corpus Christi

6300 Ocean Drive, Unit #5869

Phone 361-825-2000

peter.etnoyer@tamucc.edu

Scott C. France

Department of Biology

University of Louisiana at Lafayette

P.O. Box 42451

Lafayette, LA 70504

Phone: 337- 482-6320

FAX: 337- 482-5834

http://www.ucs.louisiana.edu/~scf4101/

Alberto Lindner

Biology Department

Duke University

Durham, NC 27708

www.duke.edu/~al18

Jose V. Lopez

Assistant Scientist

Division of Biomedical Marine Research

Harbor Branch Oceanographic Institution

5600 US 1 North, Ft Pierce FL 34946

Phone: 772-465-2400, ext 478

Fax: 772-461-2221

Email: Lopez@hboi.edu

Amy Baco-Taylor

Biology Department

MS#33, 2-14 Redfield

Wood’s Hole Oceanographic Institution

Wood’s Hole, MA 02543

Phone: 508- 289-2331

FAX: 508- 457-2134

abaco@whoi.edu

Brad Barr

Senior Policy Advisor and East Coast,

Gulf of Mexico and Great Lakes

Regional Coordinator

Office of the Director

NOAA’s National Marine Sanctuary Program

c/o US Geological Survey

384 Woods Hole Road

Woods Hole, MA 02543

Phone: 508-457-2234

Brad.Barr@noaa.gov

Sandra Brooke

Oregon Institute of Marine Biology

University of Oregon

PO Box 5389

Charleston, OR 97420

Phone: 541-888-2581

sbrooke@uoregon.edu

Andrew Bruckner

Office of Habitat Conservation

NOAA Fisheries

1315 East West Highway

Silver Spring, MD 20910

Phone: 301-713-3459

andy.bruckner@noaa.gov

Stephen D. Cairns

Research Scientist/Curator

Smithsonian Institution

P. O. Box 37012

NMNH, W-329, MRC-0163

Washington, D. C. 20013-7012

Fax: 202-357-3043

Phone: 202-786-2129

cairns.stephen@nmnh.si.edu
48

John McDonough

NOAA Office of Ocean Exploration

Expeditions Coordinator

Phone: 301-713-9444 x113

Cell: 443-994-8977

john.mcdonough@noaa.gov

Lance Morgan

Marine Conservation Biology Institute

14301 Arnold Dr., Suite 25

Glen Ellen, CA 95442 USA

Phone: 707- 938-3214

lance@mcbi.org

John Reed

Senior Scientist

Division of Biomedical Marine Research

Harbor Branch Oceanographic Institution

5600 U.S. 1 North, Fort Pierce, FL 34946

Phone- 772-465-2400 ext. 205

Fax- 772-465-2221

jreed@hboi.edu

Juan Armando Sanchez

Profesor Asistente

Departamento Ciencias Biológicas

Universidad de los Andes

Carrera 1E No 18A - 10 (J 304A)

P.O.Box 4976

Santafé de Bogotá D.C., Colombia

juansanc@uniandes.edu.co

http://www.branchingnature.org/

William W. Schroeder

Marine Science Program

The University of Alabama

Dauphin Island Sea Lab

101 Bienville Blvd.

Dauphin Island, AL 36528-4603

Phone: 251-861-7528

wschroeder@disl.org

Andrew N. Shepard

NOAA Undersea Research Center

University of North Carolina at Wilmington

5600 Marvin Moss Lane

Wilmington, NC 28409

Phone: 910-962-2446,

Fax -910-962- 2444

sheparda@uncw.edu

Les Watling

Professor of Oceanography and

Pew Fellow in Marine Conservation

Darling Marine Center

University of Maine

Walpole, ME 04573

Phone 207-563-3146 x248

Fax 207-563-3119

watling@maine.edu

Gary Williams

Department of Invertebrate Zoology & Geology

California Academy of Sciences

Golden Gate Park

San Francisco, CA 94118-4599

Phone 415-750-7080

gwilliams@CalAcademy.Org
49

Appendix 3. Deep-sea corals online

NOAA Coral Reef Information System

http://www.coris.noaa.gov/about/deep/deep.html

This portal favors Atlantic scleractinia, with links to myriad other online documents describing deep ahermatypic

reefs. It includes lovely images, and a dense bibliography on Lophelia sp. and Oculina sp.

Octocoral Research Center at the California Academy of Sciences

http://www.calacademy.org/research/izg/orc_home.html

This website created and maintained by Dr. Gary C. Williams at the California Academy of Sciences. The

bibliography from Bayer is exhaustive, at ~50 pages in its original form. The list of genera is extensive, and includes

many photographs.

Report of the International Council for the Exploration of the Sea (ICES) Study Group on Deep-sea

Corals. 2003.

http://www.ices.dk/reports/ACE/2003/SGCOR03.pdf

UN Status Report on Cold-water Coral Reefs. 2004.

http://www.unep-wcmc.org/press/cold-water-coral-reefs/

NOAA Office of Ocean Exploration Website

Gulf of Alaska http://www.oceanexplorer.noaa.gov/explorations/02alaska/welcome.html

http://www.oceanexplorer.noaa.gov/explorations/04alaska/welcome.html

The 2002 and 2004 Gulf of Alaska Seamount Expeditions (GOASEX) visited a dozen seamounts in the Northeast

Pacific Ocean with the Alvin submersible, collecting several excellent specimens. An 2002 article in the mission

background and the July 7th log describes dispersal studies. The July 14th log details some problems with

morphological classification in the Family Isididae. The 2004 August 16th log discusses octocoral sample

processing, while the August 19th log discusses and exhibits living polyp photography. August 17th describes the

“paradox of seamounts.”

North Atlantic http://www.oceanexplorer.noaa.gov/explorations/03mountains/welcome.html

http://www.oceanexplorer.noaa.gov/explorations/04mountains/welcome.html

The 2003 and 2004 Mountains in the Sea Expeditions also made some fascinating finds, and they detail

sophisticated methods in the 2003 July 15 Daily Log by Scott France. Visiting a seamount is a special event, but

returning to a previously visited seamount is truly rare. Three seamounts, Manning, Kelvin, and Bear, had

spectacular gardens of octocorals.

Gulf of Mexico http://www.oceanexplorer.noaa.gov/explorations/03mex/welcome.html

An interdisciplinary scientific team spends 12 days exploring deep-sea coral habitats in the Northern Gulf

of Mexico. Deep corals are widely distributed in a region heavily utilized by the oil and gas industry.

Hawaiian Islands http://oceanexplorer.noaa.gov/explorations/03nwhi/welcome.html

The 2003 Expedition to the Northwestern Hawaiian Islands studied the reproductive biology and

population genetics of precious corals in the Hawaiian archipelago. This site contains first ever footage of

an endangered monk seal foraging in precious coral beds.
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