NOAA Technical Memorandum NMFS-SEFSC-543 

DEMOGRAPHIC MONITORING PROTOCOLS FOR THREATENED 

CARIBBEAN ACROPORA SPP. CORALS 

BY 

Dana E. Williams, Margaret W. Miller, K. Lindsey Kramer 

U.S. DEPARTMENT OF COMMERCE 

National Oceanic and Atmospheric Administration 

National Marine Fisheries Service 

Southeast Fisheries Science Center 

75 Virginia Beach Drive, Miami, Florida 33149 

October 2006 
NOAA Technical Memorandum NMFS-SEFSC-543 

DEMOGRAPHIC MONITORING PROTOCOLS FOR THREATENED 

CARIBBEAN ACROPORA SPP. CORALS 

BY 

Dana E. Williams1,2 , Margaret W. Miller2 , K. Lindsey Kramer 1,2 

1. Cooperative Inst. of Marine and Atmospheric Science, Univ of Miami, 

4600 Rickenbacker Cswy. Miami FL 33149 

2. NOAA Fisheries, Southeast Fisheries Science Center 

75 Virginia Beach Dr. Miami FL 33149 

U.S. DEPARTMENT OF COMMERCE 

Carlos M. Gutierrez, Secretary

National Oceanic and Atmospheric Administration 

Conrad C. Lautenbacher, Jr., Administrator 

National Marine Fisheries Service 

William T. Hogarth, Assistant Administrator for Fisheries 

October, 2006 

This Technical Memorandum series is used for documentation and timely communication of preliminary 

results, interim reports, or similar special-purpose information. Although the memoranda are not subject to 

complete formal review, editorial control, or detailed editing, they are expected to reflect sound 

professional work. 

i 
NOTICE 

The National Marine Fisheries Service (NMFS) does not approve, recommend or endorse 

any proprietary product or material mentioned in this publication. No reference shall be 

made to NMFS or to this publication furnished by NMFS, in any advertising or sales 

promotion which would imply that NMFS approves, recommends or endorses any 

proprietary product or proprietary materials mentioned herein or which has as its purpose 

any intent to cause directly or indirectly the advertised product to be used or purchased 

because of this NMFS publication. 

This report should be cited as follows: 

Williams DE, Miller MW, Kramer KL (2006) Demographic Monitoring Protocols for 

Threatened Caribbean Acropora spp. Corals. NOAA Technical Memorandum 

NMFS-SEFSC-543. 91 pp. 

Author’s Addresses: 

Dana E. Williams, Southeast Fisheries Science Center, 75 Virginia Beach Dr. Miami FL 

33149 

Copies may be obtained by writing: 

The author or National Technical Information Center 

 5825 Port Royal Road 

 Springfield, VA 22161 

 (800) 553-6847 or (703) 605-6000 

http://www.ntis.gov/numbers.htm

PDF file available at https://grunt.sefsc.noaa.gov/P_QryLDS/ 

 

ii 
ACKNOWLEDGEMENTS 

We have benefited from many colleagues participation and discussion in the

development of these protocols.  Most notably, Caroline Rogers of USGS‐BRD has

labored with us in figuring out how to do this.    We are grateful for her

collaboration.    Her comments on an earlier version of this manual provided for

great improvement of clarity and description.  Financial support for development

has come from the NOAA Coral Reef Conservation Program, the National

Undersea Research Center (UNCW), and the NMFS Southeast Regional Office.

iii
TABLE OF CONTENTS 

Acknowledgements....................................................................................iii

Table of Contents....................................................................................... iv

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

Purpose ......................................................................................................1

Ecological Significance and Status of Acropora palmata ...............1

Project Goals and Objectives ................................................................2

Anticipated Products ..............................................................................3

Overview.......................................................................................................4

Main Tasks ...............................................................................................4

Optional Tasks.......................................................................................5

Materials....................................................................................................6

Personnel...................................................................................................6

Initial Survey................................................................................................8

Site Selection and Set‐up.......................................................................8

Colony Determination & Selection .....................................................9

Assessment of Tagged Colonies.........................................................11

Size ........................................................................................................11

Percent Live .........................................................................................11

Colony Conditions..............................................................................12

Healthy/Unaffected........................................................................................... 12

Recent Mortality: Rank..................................................................................... 13

Recent Mortality: Type..................................................................................... 13

Bleached Tissue: Rank...................................................................................... 18

Bleached Tissue: Type ...................................................................................... 19

Damselfish Impact ............................................................................................ 19

Skeletal Anomalies............................................................................................ 20

Overgrowth........................................................................................................ 21

Fragmentation/ Physical Damage................................................................... 22

Snail and Damselfish Census ............................................................22

iv
Corallivorous Snails.......................................................................................... 23

Threespot Damselfish....................................................................................... 24

Yellowtail Damselfish....................................................................................... 24

Photographing Colonies ......................................................................24

Mapping Acropora palmata Colonies...............................................26

Post Survey Tasks..................................................................................28

Quarterly Surveys .....................................................................................28

Optional Tasks ..........................................................................................29

Linear Branch Extension ......................................................................29

Materials & Methods ..........................................................................29

Genotype Sampling..............................................................................30

Materials & Methods ..........................................................................30

References....................................................................................................32

Glossary ......................................................................................................35

Appendices ..................................................................................................37

v
INTRODUCTION 

PURPOSE

This manual is intended to serve as a basic but evolving platform to involve

many partners in collecting comparable data sets focused on Atlantic Acropora spp.

from a variety of locations throughout the Caribbean.  This data will be utilized in

the ongoing efforts and responsibilities of NOAA‐Fisheries to assess species’ status

and promote recovery under the auspices of the US Endangered Species Act.  The

protocols described in this manual have been developed over a period of five years

of focal monitoring of primarily A. palmata in the upper Florida Keys by our team.  

We have also applied these techniques in Tortola, BVI and Curacao. Obviously,

every population and site is different and may require adaptation to the specific

monitoring methods.   

It should be noted that these techniques have been developed and applied

primarily to Acropora palmata and the remainder of the document is phrased in

terms of this species.    However, similar types of data are needed for Acropora

cervicornis and similar approaches should be appropriate, at least in theory.  In our

experience in the upper Florida Keys, A. cervicornis colonies are much more

ephemeral (i.e. they suffer a high rate of complete mortality and complete colony

dislocation) and occur at much lower density which makes following individual

colonies in statistically robust numbers impractical.  We hope to encourage partners

to apply the protocols described herein to A. cervicornis (with appropriate

adaptation) in their regions to begin to obtain similar data sets for this species.

ECOLOGICAL SIGNIFICANCE AND STATUS 

OF ACROPORA PALMATA

Acropora palmata is one of the dominant structural framework builders on

Caribbean coral reefs (Goreau 1959; Shinn 1989).    Its branched morphology and

tendency to form dense, thicket‐like, mono‐specific stands naturally provide shelter

to a variety of fish (Lirman 1999) and other reef organisms including the long‐

spined urchin, Diadema antillarum (Weil et al. 1984).   Rapid skeletal growth and

frequent fragmentation allowed it to dominate the reef crest throughout the

Caribbean (Goreau 1959) in the past.  The reef spurs found on high relief coral reefs

in the Florida Keys and throughout the Caribbean were formed in large part by A.

palmata (Shinn 1989). Because of its ability to create protective habitat, to foster reef

1
biodiversity, and to maintain the structural integrity of coral reefs, acroporid corals

are considered ecologically irreplaceable(Bruckner 2002).    

Since the 1970s acroporid species in the Caribbean have experienced extreme

and accelerating declines estimated at 90 to 98 % throughout its range (Bruckner

2002). Much of the decline has been attributed to widespread disease impacts

(Gladfelter 1982; Aronson, Precht 2001), but the overall loss is thought to have been

exacerbated by physical breakage, more frequent and intense bleaching events as

well as emerging diseases and anthropogenic effects (Porter et al. 2001).   

A multitude of stressors potentially drive A. palmata decline including toxic

substances, sedimentation and eutrophication, overfishing and related trophic

shifts, hurricanes and climate change, disease,    and possible synergistic effects.  

Presently, the ultimate cause(s) of acroporid decline and general reef demise are

difficult to parse among these various sources because they usually occur together.

Monitoring studies cannot provide robust linkages between these causes and their

specific effects.    Instead, more tractable proximal causes of decline, including

quantifiable colony conditions such as disease and bleaching, fish and snail

predation, sponge overgrowth and physical damage related to storms or direct

human interaction will be investigated, as a crucial first step in identifying the

relative contributions of specific stressors as ultimate causes of decline.  

Given the extent and ongoing nature of Acropora spp. decline, NOAA‐Fisheries

responded to a petition by the Center for Biological Diversity and the evaluation of

an expert Biological Review Team (Acropora Biological Review Team 2005) by

listing both Acropora palmata and A. cervicornis as threatened under the US

Endangered Species Act in May 2006 (NMFS 2006).  The implications of this listing

are still being worked out, but it invokes specific responsibilities for NOAA‐

Fisheries to continue to assess the status and orchestrate the recovery of these

species throughout their geographic range.

PROJECT GOALS AND OBJECTIVES

There are many studies presently in place to assess the general status and trends

of Caribbean coral reefs. Acropora palmata is often poorly represented in these

studies since its natural distribution is along the reef crest while many studies focus

survey efforts on fore reef areas. Furthermore, these studies typically survey

randomly placed transects which are not well suited to A. palmata’s presently sparse

and highly patchy distribution. As a result, very small numbers of Acropora palmata

colonies end up in the being counted in general reef monitoring studies. While this

accurately depicts the present densities of Acropora, it yields very little information

on the condition and fate of these remaining colonies. Additionally, the lack of

2
consistency of survey methods greatly hinders the ability to compare the results of

surveys between studies and, hence, regions.  

The demographic (i.e. colony‐based) monitoring approach described here aims

to determine the relative importance of the many sources of mortality for

populations of A. palmata.    This method relies on tracking the performance of

individual colonies (Fig. 1) to document the threats they face and their fate over

time. By tracking the performance of randomly selected ‘individual’ colonies, the

prevalence of a particular threat combined with the subsequent fate or lethality of

affected colonies will show the ecological relevance of the various threats. For

example, parrotfish bites may be extremely common among an acroporid

population, but if effects on a colony are minor, they may be relatively

‘unimportant’ to the viability of the population.    Management and conservation

resources can be more effectively applied based on an understanding of the relative

impacts of threats on populations.    The tracking of individual colonies results in

estimates of threat prevalence, threat impact (i.e. mortality), and change in colony

size (growth and fragmentation).  

A second aspect of this suggested monitoring approach aims to estimate

recruitment, the establishment of new colonies in a population to (hopefully) offset

the mortality from the treats observed.  This is extremely difficult to do given the

dynamic nature of fragments and the small size and likely cryptic habit of larval

settlers.  The best approach we have managed to come up with involves periodic

careful, fine scale observations (i.e. mapping) of specific plots in order to detect the

appearances of new colonies, including fragments that are attached and growing

and small crusts which may be larval recruits.

The data generated by these survey techniques aim to address the following specific

objectives:

1) Assess A. palmata density and condition in as many areas as

possible throughout its geographic range.

2) Monitor sites for changes in abundance (recruitment or

death), condition, and colony size.

3) Evaluate the relative ecological relevance and importance of

threats to A. palmata including disease, predation,

overgrowth, and physical breakage.

ANTICIPATED PRODUCTS

Survey data will yield basic population data including colony size, percent live

tissue cover, recruitment, disease prevalence, predator infestation, sponge

3
overgrowth etc. Since site and plot selection may not be random (i.e., locations

known to have Acropora spp. colonies), the described monitoring protocol is not

intended to yield regional estimates of population abundance.  However, changes

in abundance due to recruitment and mortality can be rigorously evaluated. The

random selection of colonies within the plot will yield information on the

prevalence and variability of various threats among the general population.

Ultimately, these estimates of demographic parameters such as growth, mortality,

and recruitment will be useful in future efforts at population modeling for

following and predicting population status and recovery in a rigorous quantitative

way.

OVERVIEW 

MAIN TASKS

1) Locate a site with Acropora palmata.

2) Select three 7m radius areas with moderately dense A. palmata as

study plots. If the A. palmata patch is of sufficient extent, the

location of the plots should be selected randomly. NOTE: Further

discussion and alternative means of site selection and set‐up are

included below.

3) Install a stake or other marker at the center of each plot (preferably at

random GPS point).

4) Randomly select and tag 12 A. palmata colonies using random list of

polar coordinates (see ‘Tagged Colony Selection’ field sheet,

Appendix A). DIVER B.

5) Survey tagged colonies within each plot using the ‘Initial Colony

Survey’ field sheet (Appendix A). DIVER A.

6) Photograph each colony from several angles, always including top‐

down view, with scale to document colony condition and help

identify the colony in future surveys.    DIVER A (DIVER B can

assist with locating colonies and placing scale and photoboard).

7) Careful examination of entire plot to record location of ALL Acropora

palmata (selected and unselected, particularly small crusts and

attached fragments) within each plot using the ‘Annual Plot Map’

4
(Appendix A) field sheet to enable detection of new recruits in

future surveys.

8) Take a GPS waypoint over the center stake at each plot.

9) Download and rename image files.

10) Enter field data into the database after each survey.

11) Conduct quarterly surveys to assess condition and photograph the

tagged colonies (Steps 5‐7 and 10 above).

12) Annually map all colonies in each study plot (Step 8 above).

Optional Tasks 

1) Branch linear extension (growth) measurement.

2) Tissue sampling for genotype analyses.

5
MATERIALS

Suggested materials for conducting surveys (Appendix B):  

1) Metal stakes (e.g. steel rebar 1⁄4”

(1.2cm) diam, ~14” (45cm)

length)  

2) Sledge hammer (2.5 lb (1.2 kg))

3) Aluminum numbered tags (11⁄4”

(3 cm) diam)

4) Cable ties (to fit through 7/64”

(0.28cm) hole in tag)

5) Nails, (approx 2 1⁄4ʺ  (5.7cm)

length, 1/8” (0.3 cm) diam

shank, 3/8ʺ (1 cm) diam head)  

6) Transect line (10m length) with

brass clip

7) Dive compass

8) GPS with waterproof case

9) PVC meter stick (marked 5cm)

10) Pencils (mechanical pencils

work well)

11) Rubber bands/string for

securing pencils & paper to

clipboard

12) Aluminum clipboards

13) Dura Copy Underwater paper

(http://www.riteintherain.com)

14) Photocopier or laser printer (for

printing field datasheets)

15) Underwater digital camera (>5

Mpixels & wide angle is best)  

16) 2 memory cards (bigger is better)

17) 2 camera batteries

18) Photo board (see Appendix C)

19) Field datasheet template files

(worksheet name italicized; see

Appendix A):  

a) Initial Survey.xls

i) Initial Colony Survey Diver A

ii) Tag Colony Selection Diver B  

b) Quarterly Survey.xls

i) Quarterly Colony Cond Diver

A

ii) Quarterly Sn/ Fish Census

Diver B

c) Annual plot map.xls

i) Annual Plot Map Diver A

ii) Annual Plot Map Diver B

iii) Map Diagram

PERSONNEL

This protocol is designed to be conducted by two SCUBA divers.  Since Acropora

typically occurs in very shallow habitats divers should be comfortable wearing

SCUBA gear in shallow water and should be experienced in the challenges of

6
working in surge conditions. We suggest that in general monitoring sites with less

than 2m between the TOP of the colonies and sea surface be avoided due to the

likelihood of damage to the colonies. This general recommendation can be

reconsidered based on the local conditions (e.g. very sheltered site or small widely

spaced colonies) and the experience of the divers.  Note that very shallow sites may

seem amenable to survey on snorkel, however making comprehensive colony

assessments and taking photographs is greatly compromised on snorkel.

Additionally, some tasks such as hammering (tags and stake) are impractical to

impossible on snorkel.  

Field tasks are divided such that those assigned to Diver A are those involving

relatively more training and/or subjective assessments that rely upon making

consistent observations. Diver B is assigned more supporting tasks and collects

quantitative data that is less dependent on consistent interpretation. The person

who assumes the role of Diver A should therefore be the person who is most likely

to be consistently available for follow‐up surveys. Where possible the tasks are also

distributed so that both divers will complete the survey in the same amount of time.  

7
INITIAL SURVEY 

SITE SELECTION AND SET-UP

Acropora palmata populations have variable geographic layouts in differing areas

and different reef structures that may dictate different site selection and layout (e.g.,

fore‐reef spur‐and‐groove typical of Florida Keys versus fringing A. palmata zone or

patch reefs along St. John USVI). The extremely patchy distribution of surviving

populations makes random selection of sites at the regional level impractical; any

reef site in which A. palmata is found should be considered for observation. Ideally,

each surveyed reef site should include at least three randomly‐placed, 7m radius

permanent survey plots (Fig. 2) to provide statistical replication for recruitment

assessments.  There are various situations that may dictate a less than‐ideal set‐up.

In many areas, an A. palmata patch may be too small to accommodate multiple plots

or to make random selection of plot location feasible. Any section of reef (7m

diameter) with at least 10 A. palmata colonies is suitable (Fig. 2) but areas of live A.

palmata ‘thicket’ should be avoided because the reliable designation of individual

colonies is problematic (Fig. 1). In the Florida Keys, A. palmata is generally in

remnant patches of limited extent so plots are often centered deliberately in these

small patches on adjacent or nearby reef spurs (Fig. 2). The number of field sites

selected should be based on the number that can be feasibly surveyed on a regular

quarterly schedule.    A single survey plot typically takes up to three hours to

initially tag and assess, and may take up to two hours to fully map in (both tasks

vary greatly depending on the number of colonies present). At least two people will

be required to tag colonies, and survey/map each plot and assess colony condition.   

The following list summarizes the properties of a ‘suitable plot’ location:

1) Water depth should be deep enough that divers can maneuver

over colonies easily without touching them. Specifically, the top

branches should be at least 6 feet (2m) below the sea surface.

Note that in these areas even ‘minor’ tidal fluctuations can

greatly affect the workability of a particular area. When choosing

sites it is helpful to be aware of the current tidal state.  

2) Avoid places where the colonies are growing in a thicket since

ʹdemographicʹ monitoring is necessarily dependent on being able

to distinguish individual colonies.   

3) Counting the number of colonies that are in a 7m radius area

should result in the same number (+/‐  1) if counted repeatedly.

8
This becomes more difficult when colonies are larger, more

closely spaced, or have poorly differentiated boundaries

(discussed in detail below).

To re‐iterate, the center of plot should be located randomly if feasible.  

Otherwise, after identifying a suitable plot location with an appropriate A. palmata

density that is limited in extent, a metal stake should be centrally placed using a

sledge hammer.  Care should be taken to minimize disturbance by choosing areas

for stake placement (and colony tag placement) that are not covered by coral or

sponge and have adequate surrounding working space.

After placing the center stake, the number of colonies within 7 meters of the

stake should be carefully counted by both divers.    This count should include all

colonies of any size or shape, except loose fragments (NOTE: loose fragments are

not considered in the designation of colonies for ongoing observation, but they are

counted in the plot‐level assessments of recruitment).    If there are only 10‐12

colonies within the plot, all colonies can be automatically selected and tagged.  

However, this short‐cut should be used only after observers have confirmed that

only 10‐12 colonies exist within the plot.  Keep in mind that oftentimes additional

small or remnant colonies are discovered during a careful search of the plot. For any

plot with more than 12 colonies, the colonies selected for tagging must be chosen

randomly (discussed below).   

COLONY DETERMINATION & SELECTION 

Acropora palmata, like other coral species, can often grow in unique orientations

with variable morphologies.  Colonies can fragment (after which the fragment can

become its own colony), fuse with other colonies, and suffer partial mortality which

divides a single colony into smaller isolated patches of tissue (Fig. 3).    For an

excellent discussion of these coral ‘colony ambiguities’ see, Hughes and Jackson

(1980).  For these surveys, the following rules should be applied to distinguishing

single colonies:

1) Primarily, any continuous live tissue should be considered a single 

colony (Fig. 4). 

2) If live tissue is not continuous, physically separated units of live tissue 

that are growing on the same underlying skeletal colony structure 

should be considered the same colony (Fig. 4). 

3) If live tissue is not continuous and is located on underlying 

remnant/dead structure that is difficult to distinguish (i.e., one ‘stem’ or 

two?), patches of tissue within 1 meter of each other should be 

9
considered a single colony. Patches more than 1 meter from 

surrounding tissue should be considered a separate colony (Fig. 4). 

4) If the center (based on top view) of the colony is outside of 7m it should 

not be included. 

The initial colony selection and tagging process involves two divers, referred to

from here as ‘Diver A’ & ‘Diver B’.  At this stage, Diver B is responsible for selecting

colonies and physically placing aluminum tags into the substrate and should carry

the field sheet ‘Tagged Colony Selection’ (Fig. 5, Appendix A), which lists randomly

generated coordinates (compass heading and distance from stake) to be used for

selecting colonies.    If only 10‐12 A. palmata colonies are present in the plot, all

should be tagged.    If more than 12 colonies exist in the plot, ‘Diver B’ should

randomly select colonies by following each ‘random coordinate’ listed on the field

sheet (‘Deg’ = compass heading (0‐355o), ‘Dist’ = distance from the central plot stake

(1‐7m)).  If the center of a colony lies within 0.5 m of the first random coordinate, it

should be selected and tagged.  If there is no colony at the first random coordinate,

‘Diver B’ should proceed to the second random coordinate.  Repeat this procedure

until 12 colonies have been selected.   

Each selected colony is tagged on adjacent substrate (Fig. 6).  Different kinds of

tags can be used; we use 3cm diameter, numbered, aluminum tags nailed into

adjacent bare substrate, (or if necessary, by cable‐tying it to an adjacent solid

structure) because they are relatively inconspicuous (which is important to our

permitting authorities and draws less attention of other visitors).   More apparent

tags, such as cattle ear tags can also be used.  The tag should be placed close enough

to the colony to clearly indicate to which colony it refers, but also far enough away

to prevent future overgrowth by the colony.   Less conspicuous tags, particularly,

should be placed consistently on the side of the colony between the stake and the

colony for ease of relocation.   If this is not possible, the location can be noted on

Diver B’s field sheet. The tag numbers should be recorded along with the actual

colony coordinates (heading and distance from stake) for each colony on Diver B’s

field sheet.  Tags should not be attached directly to live tissue as they can be rapidly

overgrown or irritate the surrounding tissue. Additionally, tags should not be

nailed to an already dead portion of the colony because hammering can cause distal

portions of the colony to break.  

Once a colony is tagged by ‘Diver B’, ‘Diver A’ should assess of the condition of

tagged colonies using the field sheet ‘Initial Colony Condition’ (Fig. 7, Appendix A).

The maximum length, width and height of each colony should be recorded by

‘Diver A’ (described below), along with an estimate of % live tissue, and the

presence of damselfish chimneys, bleaching, recent mortality, branch breakages,

10
predatory snails and damselfish.  A detailed description of each of these assessment

categories can be found in the section ‘Assessment of Tagged Colonies’ below.

After selecting and marking colonies, be sure to record a GPS waypoint for the

center stake of each selected plot.  It is useful to have an abbreviation for the site

and number for the plot, for example the 2nd plot at ‘French Reef’ might be ‘FR2,’

and this should be recorded on the data sheet along with the date and any other

notes that may assist subsequent surveys.  

ASSESSMENT OF TAGGED COLONIES 

Size 

On the field sheets, ‘Initial Colony Survey’ (Fig. 7) the observer will measure

colony length (‘L’), width (‘W’), and height (‘H’) in centimeters.    To ensure that

measurements are straight, its best to use a solid measuring stick or measuring tape

attached to a length of PVC pipe (or other ‘measuring stick’).  Length is a measure

of the maximum colony diameter and width should be the dimension

perpendicular to height (Fig. 8).    Height should be measured as the maximum

height of the colony from the base to top.  These measurements may include dead

portions of the colony, particularly at the base.  Importantly, height measurements

should be perpendicular to the substrate (pay close attention to the angle of the

substrate; Fig. 8).  It can be difficult to be consistent with the measurement of colony

height since parts of a colony may differ dramatically in height. Keep in mind that

this measurement is taken to estimate how much tissue is present (when used in

conjunction with the % live estimate described below) so it is more useful to

measure the height of most of the colony rather than measuring the colony height at

a single taller branch.    Even when the height is measured carefully, small

differences in where it is measured can lead to misleading differences in

measurements between surveys. Taking pictures from the sides of the colony will

allow these measurements to be roughly verified by documenting whether major

changes occurred in the colony height/structure since the previous survey.  

Percent Live 

The percentage of the measured colony that is covered with live tissue, ‘% live’

should be estimated for each tagged colony.   A colony with a score of 100% live  

(Fig. 9) would have live tissue covering the entire surface of the skeleton that was

included when length, width, and height was measured (Fig. 9, Fig. 10). It is

important to look at the undersides of branches and all sides of a colony when

11
making this estimate (Fig. 9, Fig. 10).    Remember that bleached regions of the

colony are still living and thus should be included in estimates of % live.

Colony Conditions 

The following sections describe many conditions (most are detrimental

conditions) that affect A. palmata colonies. Each colony is scored (presence/absence

or a rank estimate of severity) for each condition.   In the cases of bleaching and

recent mortality conditions the conditions are further categorized by as to ‘type’

which is recorded on the data sheet by a specific code (explained further below). For

the conditions that are scored with a qualitative rank, the rank should be scaled to

the live area of the affected colony. For all ranked conditions the following scale

applies:  

0:  Not present

1:  Present, very sparse occurrence (≤5%)

2:  Moderate colony coverage (10‐25%)

3: High coverage, condition affecting 30‐55% of the colony live area

4:  Catastrophic extent, 60‐85% of the colony’s live area affected

5:  Entire colony (≥90%) affected  

Healthy/Unaffected 

What does it look like? 

A healthy colony is defined by tissue color and a lack of other detrimental

conditions (recent mortality, bleaching, etc.). Healthy Acropora tissue (Fig. 11) varies

in color from tan to a light orange to deep orange‐brown. The intensity of the color

may vary with depth and season, but a fair range of color can be considered ‘within

normal limits’ (www.coralwatch.org uses a color chart to characterize coral health,

healthy Acropora palmata typically corresponds to E5‐E6 or D4‐D6 on their chart).  It

is important to note that tissue around the growing edges of a colony is normally

white (lacking zooxanthellae) but is not considered bleached or dead.   

How is it scored? 

Healthy colonies are designated on the field sheet only by the absence of all

other tissue conditions.  A colony may have less than 100% live tissue coverage but

may still be considered ‘healthy’ (i.e. have no noted conditions) as long as all of the

remaining tissue is normal (or within normal limits) and the dead portions are not

recently dead (see below).   

12
Recent Mortality: Rank 

What does it look like? 

When Acropora tissue dies it sloughs away leaving clean white skeleton exposed

(Fig. 12). Freshly exposed skeleton is bright white with clearly visible, sharp

corallite structure (Fig. 11b). Bare skeleton is gradually colonized by a thin turf of

algae (approximately 1 week after tissue dies) and will eventually (6‐8 weeks)

appear similar to the surrounding substrate in terms of algal colonization. Over this

time the corallite structure gradually erodes (Fig. 13).   

Dead portions of a colony are distinguished as either ‘recent mortality’ or ‘long

dead’. Recent mortality includes areas of exposed skeleton that have little or no

algal colonization and sharply defined corallite structure. These areas appear bright

white or have a light brown/green tinge once filamentous algae are present. Based

on past field observations, areas described here as ‘recent mortality’ have died

within 4 to 6 weeks. Long dead areas have heavier algal colonization including

fleshier types of algae and/or calcareous coralline algae (CCA).   Additionally, the

corallite structure gradually erodes so that long dead areas may appear completely

smooth or only slightly bumpy. There is no definitive line where an area of dead

skeleton is classified as recent or long dead (Fig. 12) and these processes vary with

season and location.  

How is it scored? 

The extent of recent mortality should be qualitatively ranked on a scale from 0‐5

(below) for each tagged colony regardless of the cause. The ranking is based on the

amount (area) of the colony affected relative to the amount of living tissue on the

colony (Figs 14‐18).  A 20cm lesion on a 30cm colony would get a much higher rank

than the same size lesion on a 100cm colony.   

0:  Not present

1:  Present, very sparse occurrence (≤5%)

2:  Moderate colony coverage (10‐25%)

3: High coverage, condition affecting 30‐55% of the colony live area

4:  Catastrophic extent, 60‐85% of the colony’s live area affected

5:  Entire colony (≥90%) affected.  

Recent Mortality: Type 

The suspected cause(s) of the recent mortality ranked in the previous step is

assigned a ‘type’ code(s) which is recorded on the field sheet (Fig. 7). The following

13
codes should be used to indicate the likely source(s) of the recent mortality that was

ranked (each described further below):  

WPx = white pox disease  

WBD = white band disease  

RTL = rapid tissue loss  

FS = snail feeding scar  

PFB = parrotfish bites  

CB= ciliate band  

CL= Cliona lesion

FP = fish poop  

Unkn = unknown cause  

In cases where there are multiple sources of recent mortality the rank should

reflect the sum of all types of recent mortality. Where possible, each type should be

recorded as primary, secondary etc. based on its relative impact to the live portion

of the colony. For example a colony with large areas of RTL and a small feeding scar

might be ranked a ‘3’ and the ‘RTL code would be recorded in the 1° column and

the FS code in the 2° column.   

For ranking purposes, all recent mortality types are grouped because the impact

on the colony is the same regardless of cause. The causes that have been identified

here are actually quite diverse including disease, predation, parasitism and

incidental causes. The purpose of distinguishing these is to identify the influence

that each has on the survivorship of Acropora colonies. The following section

describes each type of recent mortality but it is important to emphasize here that

many of the characteristic signs can be difficult to distinguish. This is particularly

true of the three types that are considered to be diseases. For this reason some

studies have grouped them as ‘acroporid white disease’. While this is a conservative

and valid approach we suggest attempting to distinguish the gross manifestations

into more specific disease types since they can easily be grouped later. It is

important to realize that rigorous ‘diagnosis’ requires identifying the pathogen

which can NOT be done visually in the field. As a result distinguishing these three

‘white diseases’ is restricted to the pattern and rate of tissue loss (Fig. 19). Tissue

loss can be band‐like, patchy or irregular and sloughy. It is easiest to interpret

disease signs when it is actively progressing (Fig. 19), but even if the progression

has halted (recrudesced) and there is light algal colonization it is possible to roughly

detect a pattern and the rate of progression.  

White Band Disease (Type Code: WBD) 

Early reports of White‐band disease (WBD) described its appearance on A.

palmata as “a sharp line of advance where the distally located zooxanthella‐bearing

14
coral tissue is cleanly and completely removed from the skeleton, leaving a sharp

white zone about 1 cm wide that grades proximally into algal successional stages”

(e.g. Fig. 20; (Gladfelter 1982)).    Reported rates of advance on Acropora palmata

average 5.5 mm d‐1 ((Gladfelter 1982)).  In addition to the relatively uniform band‐

like pattern, it also progresses in a relatively uniform manner, often radiating up

multiple branches at the same time (Fig. 20). Very close inspection of the tissue at

the ‘white band’ often has a rough or ragged appearance as though the tissue is

sloughing off of the skeleton at the ‘active disease’ front, but from more distant

inspection it typically appears very uniform.

In Florida and parts of the Caribbean it is common to see WBD only on the

undersides of branches, the progression of the disease often stops once it crosses to

the upper surface of the branch ((Williams, Miller 2006); Fig. 21).   

White Pox (Type Code:WPx) 

White pox (WPx) produces scattered patches of recent mortality (often called

‘lesions’) on A. palmata colonies. The bacteria Serratia marcescens has been identified

as the etiological agent of white pox (Patterson et al. 2002). ‘Patchy necrosis’ has also

been used (Bruckner, Bruckner 1997; Rodriguez‐Martinez et al. 2001; Weil et al.

2002) to describe lesions that are indistinguishable from those characterized as

white pox. Here we will use the term white pox (WPx) to describe multifocal,

irregular patches of recently exposed skeleton (Fig. 22).    Patches (lesions) are

typically 10‐20cm across (pers. obs.) but the size can vary, and they can remove up

to 12 cm2 (mean 2.5 cm2) of live tissue per day (Patterson et al. 2002). Unlike WBD,

this disease does not progress at a steady rate, rather, lesions seem to form rapidly

then stall (pers. obs). Once a lesion stalls and the skeleton is colonized by turf algae

the tissue margin looses its ragged appearance and no longer appears to be

‘actively’ progressing.  Stalled lesions may become active again but often they begin

to heal (Fig. 23) (Lirman 2000). Lesions are also reported to coalesce into larger

lesions (Rodriguez‐Martinez et al. 2001).  

Rapid Tissue Loss (Type Code: RTL) 

Acropora colonies are also observed with large areas of recent mortality

indicating that tissue loss occurred very rapidly, much faster and in a pattern not

consistent with the accepted descriptions of either WBD or WPx (Fig. 24).  It could

be that our present descriptions of these diseases are not adequate to describe the

full range of disease signs manifested, or it is possible that additional diseases affect

Acropora. Since this can not be determined in the field, an additional condition

named ‘rapid tissue loss’ (RTL) is used to describe colonies with large areas of

recent mortality (Fig. 24, Fig. 25).  These areas range from band‐like to lesion‐like

and with a highly ragged tissue margin.  

15
Corallivorous Snail Feeding Scars (Type Code: FS) 

The short coral snail, Coralliophila abbreviata is a common predator on Acropora

spp. and can consume more than 3 cm2/day (Miller 2001). It has a white, ridged

shell that is usually overgrown with algae (Fig. 26) making it difficult to recognize

(see also ‘Snail & Damselfish Census’ section). It resides on A. palmata colonies for

extended periods of time and is most often found in small groups (3 to 6 snails)

adjacent to live tissue (Fig. 26) but may wander to other portions of the colony when

‘resting’ (not feeding; Fig. 26).  

Feeding scars vary in size from a few square centimeters to 100 cm2 depending

on the size and number of snails feeding in that area. The shape of a feeding scar is

highly variable, but often has a scalloped margin and is almost always found along

the tissue margin of a colony leading into a ‘long dead’ portion of the colony. Snails

typically begin feeding at the base of a colony and leave behind a path (Fig. 27) of

dead skeleton that can be obvious on larger colonies.

Feeding scars can be distinguished from disease associated recent mortality first

and foremost by the presence of snails (see section ’multiple types’ for exceptions).

Additionally, while the overall shape of the scar may be irregular, the actual margin

of live tissue is typically relatively clean compared to ‘active disease’ areas. Snails

are only rarely observed feeding in an area surrounded by live tissue (Fig. 28), thus

while it is conceivable that they could produce a WPx type lesion, it is unlikely that

they would produce several lesions on one colony and then disappear from the

colony.   

However, feeding scars are often found on the undersides of branches and can

appear similar to WBD (Fig. 28).    In these cases, presence of the band of recent

mortality on multiple branches is more consistent with WBD.  If thorough searching

of the colony does not reveal any snails and the recent mortality is otherwise

consistent with WBD then it should be categorized as WBD.  

Rapid tissue loss has been observed to coincide with snail feeding activity, when

snails are present this can only be presumed if the area of recent mortality is

significantly larger than can be explained by the number of snails present, or if the

margin is highly irregular and ‘ragged’. If the observer is in doubt and snails are

present and no other disease lesions are present on the remainder of the colony,

then the recent mortality should be presumed to be a feeding scar.    

Parrotfish Bites (Type Code: PFB) 

Parrotfish typically feed by scraping reef substrate for the endolithic algae and

sponges. However scattered scrape marks (Fig. 29) are not uncommon on live A.

palmata.  They are distinctive from other types of recent mortality by the absence of

corallites and chiseled appearance of the skeleton (Fig. 29). Stoplight parrotfish

16
(Sparisoma viride), are known to form loose feeding aggregations and repeatedly

scrape a cluster of coral heads over days to weeks leaving scars up to 200cm2 on

Montastraea annularis (Frydl 1979). This ‘focused biting’ behavior (Humann,

DeLoach 2002a; Bruckner et al. 2003) is poorly documented in A. palmata but clearly

occurs sporadically (Fig. 30).    This natural but destructive feeding habit removes

live tissue and abrades the skeleton, occasionally affecting large continuous

portions of a colony, typically along the entire margin of large branches.  

Parrotfish bites should only be recorded if they affect the colony’s live tissue,

when they occur on long dead portions of the colony they do not need to be

recorded. Additionally, a colony may bear marks that resemble parrotfish bites

from a distance, however on closer inspection the corallites and polyps/tissue are

intact, and the marks are simply bleached (Fig. 31). This is likely to be a stage of

recovery from parrotfish bites (pers obs) but further documentation is needed.  

Regardless of the explanation, these bleached areas are not recent mortality and

thus should not be included in the rank, instead they will be included as bleached

tissue (discussed below).  

Ciliate Band (Type Code: CB) 

A dark greenish‐black band associated with areas of recent mortality (Fig. 32)

has recently been identified as infestation by the ciliate Halofolliculina sp. (Croquer

et al. 2006). It is uncertain whether or not the ciliates are responsible for the recent

mortality or if they are opportunistic colonizers of already compromised tissue.

Regardless they are included here as a ‘type’ of recent mortality. Based on

observations in Florida, when ciliates are present, they are generally the secondary

type of recent mortality (Fig. 33) as they are often observed associated with other

disease signs, feeding scars and Cliona lesions (discussed below). While it is

altogether possible that they alone could cause tissue mortality, they should only be

considered primary if they are present in all instances (i.e. lesions) of disease on that

colony.   

Cliona Lesions (Type Code: CL) 

Cliona laticavicola is an orange excavating sponge found commonly in reef

substrate. It does not form a thallus, but exists only inside substrates. It is visible

only where its ostia burrow to the surface (Fig. 34) or in freshly fragmented

substrates.  

Cliona sp. can grow aggressively around the A. palmata’s live tissue margin and

appear to drive tissue retreat.   In cases where sponge is associated with a lesion

(recent mortality), it should be included in the ranking and noted as type CL. Cliona

present on the long dead portions of the colony or in healing lesions should not be

assessed as recent mortality.   

17
Fish Poop Lesions (Type Code: FP) 

The flat palmate structure of A. palmata branches allows fish fecal material (Fig.

35; fish poop!) to settle and remain on the branches, particularly in calm weather. In

many cases the material will smother the tissue underneath and result in a lesion

((Weil et al. 2002; Williams, Miller 2006)).  Acropora palmata colonies are commonly

seen with a few scattered small (<10cm) lesions.  These can appear similar to WPx

lesions and in fact it has been suggested that they are related (Bruckner 2002).  

Lesions caused by fish poop are often crescent shaped (Williams, Miller 2006) with

small amounts of ‘poop’ remaining in the lesion.    These types of lesions heal

rapidly.   

While fish poop is common on colonies, it does not always result in a lesion and

should only be recorded as a ‘type’ of ‘recent mortality’ if it is responsible for a

notable portion of the recent mortality ranking.  

Unknown (Unkn) 

If recent mortality is present and can not be identified as one of the described

types, it should be recorded as unknown with notes as to likely types. In these cases

taking several photographs (both wide angle and close up; see section

‘Photographs’ below) will allow it to be re‐evaluated later. When categorized as

‘unknown’ the surveyor should note on the field sheet any additional information

that may help evaluate it later. It is particularly important to note whether the tissue

is bleached or if it is gone (i.e. recent mortality), while this can be difficult to

determine in the field it is even harder to determine from a photograph (white areas

do not photograph well). In many cases photographs taken at the next survey will

show whether the area was dead or just bleached.  

Bleached Tissue: Rank 

What does it look like? 

Acroporid tissue changes color from healthy to pale to completely translucent,

bleached tissue when zooxanthellae are lost (Fig. 36).  Totally bleached portions of

colonies are bright white and from a distance look similar to very recent mortality.

However, on close inspection polyps are visible and the sharp corallite structure of

the skeleton is obscured by the translucent tissue. Bleaching may affect whole

colonies in extreme situations, but more commonly, only part of the colony is

bleached (Fig. 37). Furthermore, the affected tissue itself may be less than

completely bleached resulting in pale tissue.    

Tissue bleaching occurs for a variety of reasons including thermal stress,

changes in light, tissue trauma or other acute stresses. Mass bleaching events are an

18
extreme example of acute bleaching, apart from these events, whole colonies are

rarely fully bleached. Partially bleached colonies typically recover before the next

survey (3 months).  In contrast there are colonies with discrete patches of bleached

tissue that persist for long periods (years). These patches are most often triangular

or wedge shaped and found around the base of the colony (Fig. 38). Since there are

no published descriptions of these patches we will refer to them as ‘Wedgies’.  

Although they seem completely benign, their cause is uncertain and by noting their

occurrence we can document any effects and perhaps eventually attribute causes.  

How is it scored? 

Bleaching is ranked on a scale from 0 to 5 (see scale in ‘recent mortality’ section

above) on the field sheet.    Pale tissue is treated here as a less severe form of

bleaching. The ranking should reflect both the area bleached/pale tissue and the

degree to which it is bleached relative to the amount of live tissue on the colony. As

with the recent mortality rank, a small patch of bleached tissue would get a lower

rank on a large colony than would the same sized patch on a smaller colony.

Bleached Tissue: Type 

Codes should similarly be applied to describing the nature of colony bleaching.

Bl= bleached (intact, colorless live tissue)

W= “wedgies” (small, wedge‐shaped areas of bleached live tissue)

R= rings (white ring‐shaped areas of live tissue)

All bleaching should be recorded as the type code ‘Bl’ regardless of severity with

two exceptions. Two specific patterns are coded separately due to their distinctive

appearance. First, are the afore mentioned ‘wedgies’ They are typically small

(~10cm max length) and found at the margins of a colony though they can occur

anywhere on the colony. They are observed to persist for long periods (e.g., >2yrs of

observation in the Florida Keys) and do not seem to change or adversely affect the

colony. They are recorded as ‘wedgies’ (W) under bleaching so that an estimate of

their prevalence and distribution can be documented. Second, rings of bleached

tissue (Fig. 39) are sporadically observed on colonies. These are thought to result

from yellowtail damselfish bites (Bruckner et al. 2003). The bleached tissue regains

it’s normal color within weeks, and frequently new rings will appear on the colony

in that time.   

Damselfish Impact 

What does it look like? 

19
Threespot damselfish (Stegastes planifrons; referred to from here simply as

‘damselfish’) can aggressively establish territories (Fig. 40) on large areas of A.

palmata (Robertson et al. 1981). They are thought to farm algae ‘lawns’ (Fig. 41)

within that defended territory. Impact on live tissue in a territory ranges from

sparse bites that heal into chimney‐like structures (Fig. 40) to lawns which result

from dense clusters of bites that are rapidly colonized by algae leaving no live tissue

on that portion of the colony. Bites initially appear as small (~1cm) spots of bleached

or recently dead tissue usually interspersed with chimneys or other signs of

damselfish activity. Damselfish nests (Fig. 42) also impact A. palmata colonies; they

prefer to nest on the undersides of branches or sheltered portions of the colony.

Eggs are deposited in the nests year round but seem to be more common in the

summer. When eggs are present in the nest, they appear as dark purple‐black

masses, but when vacant, these areas of dead skeleton can resemble white‐band

diseased areas or healing lesions (Fig. 42). The same nest is used repeatedly which

reduces the impact on live tissue, but when new nests are built live tissue is often

cleared to create the nests.

How is it scored? 

The amount of perceived impact (chimneys, bites, lawns & nests) to the colony

should be ranked from 0 to 5 (Fig. 43 Fig. 44 Fig. 45) on the field sheet. Since these

bites can include both bleached and dead areas that are too small to accurately

classify and/or rank as either condition, they are included in the damselfish impact

rank.    Lawns are difficult to distinguish from other long dead portions of the

colony, these should only be included in the rank if the area is surrounded with

chimneys or other evident damselfish‐ inflicted damage.  

Skeletal Anomalies 

What does it look like? 

Several types of skeletal/growth anomalies can affect A. palmata colonies.  While

the cause is unknown, tracking them over time will allow us to document their

impact on overall colony performance.

Neoplasia (Np) (Fig. 46) are smooth bulbous growths on the colony surface with

little or no coloration.    Neoplastic regions typically occur along the new growth

edge of A. palmata colonies.  More technical description can be found in (Peters et al.

1986; Bruckner 2003; Gateno et al. 2003). Hyperplasia (Hp) (Fig. 46) is another

condition commonly observed and appears as polyp calices that display accelerated

and irregular growth. Affected polyps are large and pronounced and protrude

above the colony surface (Bruckner 2003).   

20
Colonies often have area around the margin where the polyps (corallites) are

densely spaced and lacking zooxanthellae (Fig. 47). The cause of this pattern is

unknown but often Yellowtail damselfish are closely associated so it is possible that

it is territory or nesting related. For the purposes of this study, this phenomenon is

called dense white polyps (DWP) and is broadly categorized as a skeletal anomaly.

Additional cases of irregular corallite structure are also observed and simply

classified as ‘unknown’. Note that corallite structure can vary somewhat between

colonies but this category is reserved for cases where the skeleton on a portion of a

colony is notably different from the remainder of the colony.   

How is it scored? 

Skeletal anomalies are recorded only by the type of anomaly without regard to the

severity of the anomaly. Skeletal anomaly types are recorded on the field sheet with

the following codes:

Np = neoplasia

Hp = hyperplasia

DWP = dense white polyps

Unkn = unknown   

Overgrowth 

What does it look like? 

Competition for space often occurs between sessile organisms, Acropora included

(Fig. 48, Fig. 49). Its fast growth rate gives it an advantage in many situations, but

the competitive interaction probably slows growth at best, but it often results in

retreat of the tissue margin. This is especially true with Cliona langae (Fig. 48), an

encrusting sponge (Weil et al. 2002; Bruckner et al. 2003) that is found in the Greater

Antilles portion of the Caribbean.   In Florida, overgrowth most often occurs with

Erythropodium caribaeorum (Fig. 49), Millepora complanata, colonial tunicates and

macrosponges. Regardless of the competing species, the interaction between

Acropora and a competitor is typically marked by a margin of blanched tissue (pers.

obs.).

How is it scored? 

This sponge and other overgrowing organisms are not typically associated with

recent mortality so it is not included as a ‘type’ of recent mortality. Instead it is

classified as overgrowth and the type of organism should be recorded on the field

sheet.  

21
Fragmentation/ Physical Damage 

The natural life history of acroporid corals relies strongly on fragmentation and

the successful reattachment and growth of the fragments (Fong, Lirman 1995). This

reproductive strategy has associated costs and benefits. It can foster rapid

proliferation and spatial dominance; however it can also result in losses since loose

fragments are more vulnerable to export or physical stress. Once broken, branches

may nestle into surrounding reef structures and reattach, or they may be

transported to sandy areas where they become abraded or smothered and can not

attach. Although, asexual reproduction through fragmentation is a natural process,

fragments are less likely to be retained in lower density stands (Fong, Lirman 1995;

Miller et al. 2002), pers. obs.) typical of present populations.   

What does it look like? 

Branch breaks that occurred recently are fairly obvious, they are bright white,

smooth and angular (Fig. 50). These areas are colonized by algae in the same way as

recent mortality but should not be included as recent mortality since the tissue did

not die. Older breaks can be difficult to recognize in the field, but can be detected in

the photo series for each colony.    

How is it scored? 

Obvious broken branches on each colony should be counted and recorded on

the field sheet according to the approximate diameter of the break (it is not

important to take precise measurements of each break encountered).  

sm = small <5cm  

med = medium 5cm‐10cm  

lg = large 10cm‐20cm  

xlg = extra‐large >20cm

Extreme disturbance events may ‘fragment’ entire colonies rather than

individual branches.  This would be scored as an extra‐large break.  

Snail and Damselfish Census 

In addition to characterizing the A. palmata colony itself, we also document the

associated predators.   On each tagged colony, the number of corallivorous snails

(Coralliophila abbreviata (Fig. 51) and C. caribbaea, Threespot damselfish (Stegastes

planifrons) (Fig. 52), and Yellowtail damselfish (Microspathodon chrysurus) (Fig. 53)

associated with each colony should be recorded on the field sheet.  

22
Corallivorous Snails 

Several types of snails are commonly found on A. palmata colonies, however

only two species of Coralliophila spp. (Fig. 51) are significant predators of live tissue,

and only these are to be counted in surveys. Coralliophila abbreviata (yellow

operculum) is by far the most common species, but occasionally Coralliophila

caribbaea (purple‐brown operculum) is found feeding on Acropora colonies. They can

be difficult to distinguish without removing them from the colony, so for the

purposes of these surveys, both species can be counted and recorded under ‘snails’

on the datasheet.     

Other snails (especially Thais deltoidea; Fig. 51) are commonly found on Acropora

colonies.  While the clean shells of these two are very different, both are very well

camouflaged by heavy algal encrusted growth on their shells and may be difficult

to distinguish while resting on the colony. When removed from the colony they are

readily distinguished by the color of their operculum and body: Coralliophila

abbreviata are bright yellow while T. deltoidea are deep purple. However, past

observations have shown that attempting to replace snails that are removed from a

colony for identification often fails. Frequently they are on undersides of branches

and can’t be replaced, and replacing them in other areas at the very least changes

the placement of the feeding scars, but often the snail will wander off of the colony.

All of these outcomes will affect the data being collected so should be avoided

among tagged colonies. Thus, surveyors are encouraged to develop a search image

by searching for snails on Acropora colonies outside of the study plot where they can

be removed for closer inspection.   

There are other visual ways to distinguish these snails that do not require

disturbing them. First: Snails found associated with a feeding scar are most likely

Coralliophila spp.    Thais deltoidea does not feed on coral tissue, so is only

coincidentally observed in the vicinity of a feeding scar. However, Coralliophila

abbreviata is occasionally found with no apparent feeding scar, so the inverse is not

necessarily true. Second: Thais deltoidea is typically observed alone (or occasionally

feeding on another snail) while C. abbreviata is typically found in groups of 2 or

more (most often 3 to 6).  

Third: In spite of heavy algal growth it is often possible to see the rounded spires

on T. deltoidea. C. abbreviata does not have this knobby appearance and it is

occasionally possible to see the fine ridges through the algal growth.  

Coralliophila shells that are heavily overgrown by algae can also appear similar to

clumps of algae, tunicates or other non‐descript algal encrusted lumps.  Wiggling

the lumps very gently can reveal if they feel fixed (i.e. skeletal substrate) or ‘give’

slightly (a snail shell wobbling on an attached foot), or have a more mushy texture

such as a tunicate.

23
Based on thorough visual inspection and touch verification for suspicious algal

covered lumps around the margins of live tissue, record the total number of

corallivorous snails for each tagged colony.

Threespot Damselfish 

Threespot damselfish (Stegastes planifrons; Fig. 52) are commonly associated with

Acropora spp. and other corals. They aggressively defend a localized territory on live

coral colonies and can inflict substantial damage to the live tissue in their territories.

In addition to assessing the chimneys and other damage associated with their

territory (described above), the number of adult (‘Ad’) and juvenile (‘Juv’) fish

present on a colony are recorded during each survey. When colonies are closely

spaced, a single fish may be defending a territory that spans across portions of 2

separate colonies. NOTE: In these cases the one fish should be counted twice (and

note that it was the same fish), once for each colony since both colonies are being

affected. Juveniles can be highly abundant but do not have the same impact as do

the adults. These should be recorded in the separate column labeled ‘juv’.

Other species of damselfish similar in appearance are often associated with A.

palmata, and can even be more abundant than the threespot, but do not seem to

result in colony damage so are not included in these surveys.  In particular, Dusky,

Beaugregory and Cocoa damselfishes can be common and similar in appearance to

threespots (as adults). For specific distinguishing traits see one of the numerous

field guides to reef fishes (Humann, DeLoach 2002b).  

Yellowtail Damselfish 

Yellowtail damselfish (Microspathodon chrysurus; Fig. 53) are also associated with

Acropora palmata colonies and frequently overlap with the territory of a threespot,

dusky or other damselfish. Adults typically roam over a larger area than the

threespot (often multiple colonies) while juveniles remain in more confined areas.  

Their effects on the colony are both less dramatic and distinctive. Their bites may

produce small (2cm) circular bleached patches ((Bruckner et al. 2003)) and they also

occasionally nest around live Acropora palmata tissue. The number of adult and

juvenile yellowtail roaming over a colony should be recorded at each survey. It can

be difficult to determine the ‘territory’ of an adult yellowtail but if an individual

appears closely linked to the colony it should be counted.  

PHOTOGRAPHING COLONIES

24
Photographs are used to document what was observed during a survey so that

future comparisons can be made. The data collected during the survey is needed for

quantitative analyses, and could be adequate alone, however many of the

observations in the field are qualitative and subjective so that taking photographs

can allow data to be verified for consistency, particularly when comparing between

observers.  

At each survey, all tagged colonies should be digitally photographed from

several angles with scale and colony tag number included. Diver A should take the

photographs since it is important to document the conditions noted in the colony

condition assessment. At minimum, photographs should be taken from each of

these angles:  

1) Planar (top down) view of the whole colony with scale (meter

stick) and tag number (on ‘photoboard’ see below and

Appendix C).

2) Side view of the whole colony from at least 2 sides, but

preferably more for larger colonies).

3) Close‐up view of any special conditions affecting the colony

(disease lesions, predation feeding scars, etc).

Tips:

1) See the suggested ‘photoboard’ (Appendix C) for a method of

including the colony tag number in the photograph which

is stable (even in moderate surge). This board can also serve

as scale for smaller (<50cm) colonies.

2) The scale/photoboard should be placed on top of the colony

and level for the planar view photograph. It will only work

as accurate scale if it is parallel to the camera lens and

colony surface. If placed and removed gently it will not

damage the coral, however it is preferable to place it on

already dead areas, if present.  

3) In cases where the colony is large and/or too shallow for the

planar view to fit in one frame, simply take several

overlapping photos from far away as possible. In these

cases it is still useful to have an additional view with the

entire colony in one frame though it will be taken from an

oblique angle.  

4) If the scale or photoboard obscures important colony details,

a second picture can be taken without them.  

25
5) The photoboard and scale only need to be in the first few

whole colony photos so long as all photos of that colony are

taken consecutively and the board is placed in the first

photo of the next colony.  

6) We have found it to be more efficient to photograph all

colonies after the colony assessment data are collected.  

7) If Diver B has completed all tasks, he/she can assist Diver A

by locating and the next colony, placing the scale(s) and

preparing the photoboard with the proper colony

designation while Diver A completes the close‐up condition

photos for the previous one.  This allows the photographer

to concentrate on the colony at hand and saves time.  

8) If many cases of the same type of condition (e.g., many

disease lesions) are present on one colony it is not necessary

to photograph each one, select a few representative shots.  

A typical plot with 12 colonies will have approximately 6 photos per colony

meaning 72 files per plot, so it is crucial to keep the files organized.  At the end of

the day, the pictures should be downloaded and renamed using a conventional

naming scheme. Our surveys use the following mask: SI###‐1a where SI is a 2 letter

site abbreviation and ### is the three digit tag number followed by a dash then the

survey number (1 in the above example) and a different serial letter assigned to

each image. This task is a lot easier using a ‘batch rename’ tool available in many

image processing software programs or as standalone shareware programs

available online.  The date and time and other file information will be retained as

part of the file information so it is not necessary to include it in the file name.  

MAPPING ACROPORA PALMATA COLONIES 

Use the field sheets ‘Annual Plot Map’ to record the coordinates (distance from

stake and compass bearing) of all A. palmata colonies within each plot, including

tagged and untagged colonies, very small colonies and attached fragments. Loose

fragments that are not mapped should be tallied as they are encountered in the

mapping process. Use the rules described above to distinguish the individual

colonies to be mapped.    Plot mapping will require at least two divers, and will

likely require at least 1 long dive per plot.  

26
Prior to mapping, tag numbers and coordinate information (‘distance from

stake’, ‘compass heading from stake’) for tagged colonies should be recorded on the

field sheets ‘Annual Plot Map’, then confirmed during the mapping process.   

During the mapping dive, Diver B should remain over the central stake, and

hold the measuring tape in place while Diver A extends the measuring tape to the

center of each colony (Fig. 54).  ‘Diver B should read the compass heading from the

central stake to the colony and signal it to Diver A.   IMPORTANT: Note that the

compass should be held at least 50cm from the stake to avoid magnetic distortion

from the stake.  Diver A should assign sequential numbers to each untagged colony,

and record the distance from the central stake to the center of each colony, the size

(length, width, and height), % live tissue and colony type for each colony on the

field sheet ‘Annual Plot Map’.   This process should be repeated for all A. palmata

colonies within a plot, including very small colonies and attached fragments.   As

the diver‐team comes across a tagged colony, the original coordinates (determined

during the tagging process) can be quickly confirmed.   

It is worth a specific reminder that dive compasses are designed to be read

through the side window, not from the top.  Many experienced divers tend to forget

or (worse) be inconsistent in reading their compass from the top.    MAKE SURE

YOU ARE CONSISTENT both in reading your compass through the side window

and in the direction the heading of each colony is recorded (i.e. FROM stake TO

colony).   By the same token, if on subsequent annual mapping surveys you have

trouble locating many or all of the previously mapped colonies, it might be worth

checking the opposite heading.

While assigning ‘colony type’ for each colony on the data sheet (Diver A), avoid

touching the colonies or fragments as the re‐attachment process can be easily

interrupted.  Try to assess the nature of each fragment visually only!   

On the data sheet, ‘type’ should be filled in as follows:

• Branched colony (BC)  ‐  ‘normal’ looking colony with branches, may

have some partial mortality.

• Remnant colony (RC) ‐ live tissue mostly encrusting, no/few branches .

• Attached fragment (AF)  ‐ live fragment (usually a branch) with some

signs of attachment to the reef.  If attachment of a fragment can not

be determined visually, map it and note the uncertainty. It is

important not to touch fragments since they may be in the early

stages of forming and attachment.  

• Stable Fragment (SF) ‐ Occasionally large (>75cm) portions of colonies

are broken off and found ‘loose’ in the plot. While they may not be

attached to the substrate they can be considered ‘stable’ due to their

weight, structure and/or location. These ‘fragments’ should be

27
counted among the colonies initially chosen for tagging and

mapped along with other attached colonies.  

• Loose Fragment (LF) ‐ live fragment (usually a branch), loose in rubble

pile or on reef, obvious fresh white break. These should be tallied

during mapping but do not need to be mapped or measured. Note

that occasionally loose fragments land on living tissue, these should

not be considered separate fragments, but rather part of the colony

on which they have landed.

POST SURVEY TASKS

1) Download and rename photo files as described above under

‘Photographing Colonies’; organize files by colony (i.e., one

folder per colony).  

2) It is worthwhile to compare the previous survey photo of each

colony along with the most recent to verify that the colony was

correctly identified and that the colony condition was accurately

recorded.  

3) Make a contact sheet with one picture of each colony to assist

locating colonies on the next survey. Many image processing

software programs will do this automatically!.

QUARTERLY SURVEYS 

Subsequent surveys should ideally occur quarterly (approximately every three

months) or sooner in the case of a known disturbance event (e.g. mass bleaching or

hurricane).   Tasks 5 and 6 are repeated during quarterly surveys.   However both

Diver A and B will collect data since Diver B is not occupied with colony selection.

Diver A will use the ‘Quarterly colony condition assessment’ field sheet (Fig. 55) to

record colony condition observations. Diver B will locate the tags and clear them so

that the number can be read. Then Diver B will count the snails and damselfish

using the ‘Quarterly snail and fish census’ field sheet (Fig. 55).    If an additional

diver is available to participate, it would be desirable for that person to perform

branch extension measurements/photos (see optional tasks).

The small aluminum tags can be difficult to locate, even after only 3 months.

Diver B should be prepared to scrape turf algae or chip CCA from the tags so that

they can be read. In case some tags can not be found, contact sheets made prior the

survey can greatly improve the chances of relocating a colony.    Furthermore, in

28
cases of ambiguous colony boundaries, these photos can improve the reliability of

colony measurements.   

OPTIONAL TASKS 

LINEAR BRANCH EXTENSION

Overall colony dimensions and % live tissue give one measure of how much

colonies (or total live area) are changing over time (growing or shrinking), but these

measures are imprecise.  We would also like to measure the rate of branch extension

as a different, more precise measure of colony growth.  This is also a measurement

that has been used in previous published studies of Acropora palmata ecology and

physiology and so can be compared to historical (e.g. pre‐die‐off conditions).

Materials & Methods 

1) Small, thin, beaded cable ties, preferably at least two colors (e.g., 

http://www.coleparmer.com/catalog/product_view.asp?sku=0683050) 

2) Small, cm-scale ruler 

3) Digital underwater camera 

We have been marking ~ 4 branches per colony and will use these

measurements to calculate the average linear extension for the colony.  A cable‐tie

loop is placed tightly around (preferably) each of these four branches to form a

benchmark. The small (~5 in), flexible beaded cable ties are best for this application

as they are less obtrusive and the beaded flexible shaft of the tie will nestle and

“grip” amongst the individual corallites on the branch whereas heftier cable ties

would crush more corallites in order to get a firm fit. More than one cable tie can be

linked together if needed.  It is important that the cable tie be tight and not move

since its location will be used to gage branch length.  Place the cable tie fairly close

to the branch tip (1‐2 cm) to improve the resolution of the length measurements.

Note that this cable tie will likely be completely overgrown within ~6 months so

sequential cable ties will need to be used to monitor branch growth over time.

We have devised a designation system with the branch nearest the plot stake

being designated “A”, and then working clockwise around the colony designating

branches “B”, “C”, and “D”.   A top down photo is taken of the whole colony to

facilitate locating the designated branches on future surveys.  The small cable ties

are visible in a full colony photo at the initial placement, but may be overgrown by

29
coral tissue on subsequent surveys, so the full colony photo can be used to focus

ones’ search for the small, loose tail of the cable tie.   

At the initial marking and at subsequent surveys, the maximum distance from

the cable tie to the branch tip should be recorded.    Depending on your capacity

(field vs. lab time), this can be done in the field (most effectively using a transparent

ruler), or from measuring close‐up digital photos of each branch tip, including an

opaque ruler next to the branch in the photo.    Linear extension (growth) is

calculated as the difference in length between subsequent surveys.

GENOTYPE SAMPLING 

Acropora spp. are highly clonal organisms.  That is, many colonies originate from

fragments of mother colonies (i.e. branch that breaks off and re‐attaches to the reef

to form a new colony) rather than from sexual larvae.  Hence, many colonies within

a site may be genetically identical.  We hope to learn two things from genotyping

the monitored colonies; 1) the genotype of a colony may have a strong influence on

colony characteristics and performance (e.g. growth rate, disease susceptibility) that

we are monitoring and we want to understand this relationship better, 2) we want

to know relatively how many genotypes are present in each plot as this gives an

indication of the relative success of larval recruitment in that population.

Note: The tissue sampling described here requires extensive permitting, special

shipping considerations, and specialized laboratory analysis.  Please contact SEFSC

for advice and coordination prior to undertaking genotype tissue sampling.

Materials & Methods 

1) Snipping tool (e.g. garden pruners or wire cutters and/or small chisel 

(1/4 inch) and hammer 

2) Small plastic bags with labels (e.g. 60ml size Whirl-pak) 

3) Cryo-vials 2 ml for sample storage 

4) Ethanol (non-denatured > 95%) for sample preservation 

5) Rite-in-the-Rain paper (http://www.riteintherain.com/ and search for 

‘grid sheets’) for interior labels in EtOH. (Note that plastic types 

waterproof paper such as Dura-Copy will dissolve in ethanol!) 

The goal is to obtain a very small amount of live Acropora spp. tissue with the

minimum harm/disturbance to the colony.  As little as 0.5 cm2 of tissue is adequate

to extract enough DNA for the analysis. We have found the most precise way to

obtain such small samples of A. palmata is using a flat‐mouthed snipper (as you

30
would use for snipping off nail heads) to snip a tiny bit of branch tip or the smallest

size of wood‐chisel (1/4 inch) to chip a small piece of tissue/skeleton from the

encrusting base of a branchless colony.    Snipping branch tips is probably more

feasible for A. cervicornis since they are less likely to have an encrusting base.  Place

this small sample in a plastic bag in the field with a label with the colony number.  

You can either pre‐label the bags, or include a small label inside the bag.  On the

surface or upon return to land, the sample should be transferred from the bag into a

small cryo‐vial (or larger sample jar if needed).  Forceps can be helpful in this step

and the sample can be further fragmented if needed to fit into the small vials.  

Construct a small label on the Rite‐in‐the‐Rain paper with pencil to fit inside the

vial and include the colony number, site/plot number, and date.  It is also good to

label the outside of the vial with the same information but this is not sufficient as

the ethanol can dissolve permanent marker.  Inside labels are NECESSARY.

If possible, it is good to keep the samples in a freezer (‐80oC freezer is ideal) until

shipment and analysis can be arranged.    Again, prior arrangements need to be

made for shipping and analysis of these samples.

31
REFERENCES 

Acropora Biological Review Team (2005) Atlantic Acropora Status Review Document. Report

to National Marine Fisheries Service, Southeast Regional Office.

Aronson RB, Precht WF (2001) White‐band disease and the changing face of Caribbean

coral reefs. Hydrobiologia 460:25‐38

Bruckner AW (2003) Field Guide to Western Atlantic Coral Diseases and Other Causes of

Coral Mortality. In: E. Corcoran EG (ed). UNEP‐WCMC, Marine and Coastal

Programme, Cambridge, UK  

Bruckner AW (2002) Proceedings of the Caribbean Acropora Workshop potential application

of the U.S. Endangered Species Act as a conservation strategy: April 16‐18, 2002, Miami,

Florida NOAA Technical Memorandum NMFS‐OPR‐24, Silver Spring, MD, 184 pp.

Bruckner AW, Bruckner RJ (1997) Outbreak of coral disease in Puerto Rico. Coral Reefs

16:260‐260

Bruckner AW, Corcoran E, Green E, OʹCaroll J (2003) Field guide to western Atlantic coral

diseases and other causes of coral mortality. UNEP World Conservation Monitoring

Centre; Marine and Coastal Programme, Cambridge, UK

Croquer A, Bastidas C, Lipscomp D, Rodriguez‐Martinez RE, Jordan‐Dahlgren E, Guzman

HM (2006) First report of folliculinid cilliates affecting Caribbean scleractinian corals.

Coral Reefs 25:187‐191

Fong P, Lirman D (1995) Hurricanes cause population expansion of the branching coral

Acropora palmata (Scleractinia): Wound healing and growth patterns of asexual recruits.

Marine Ecology 16:317‐335

Frydl P, Phillips RC, McRoy CP (1979) The effect of parrotfish (Scaridae) on coral in

Barbados, W.I. Int. Rev. Gesamt. Hydrobiol. 64(6):737‐748  

Gateno D, Leon A, Barki Y, Cortes J, Rinkevich B (2003) Skeletal tumor formations in the

massive coral Pavona clavus. Marine Ecology‐Progress Series 258:97‐108

Gladfelter WB (1982) White‐Band Disease in Acropora palmata : Implications for the

Structure and Growth of Shallow Reefs. Bulletin of Marine Science 32:639‐643

Goreau TF (1959) The ecology of Jamaican coral reefs: I. species composition and zonation.

Ecology 40:67‐90

32
Humann P, DeLoach N (2002a) Reef Coral Identification : Florida, Caribbean, Bahamas :

including marine plants. New World Publications, Jacksonville, Fla

Humann P, DeLoach N (2002b) Reef Fish Identification : Florida, Caribbean, Bahamas. New

World Publications, Jacksonville, Fla

Lirman D (2000) Lesion regeneration in the branching coral Acropora palmata: effects of

colonization, colony size, lesion size, and lesion shape. Marine Ecology‐Progress Series

197:209‐215

Lirman D (1999) Reef fish communities associated with Acropora palmata: Relationships to

benthic attributes. Bulletin of Marine Science 65:235‐252

Miller MW (2001) Corallivorous snail removal: evaluation of impact on Acropora palmata.

Coral Reefs 19:293‐295

Miller MW, Baums IB, Williams DE, Szmant AM (2002) Status of Candidate coral, Acropora

palmata, and its snail predator in the upper Florida Keys National Marine Sanctuary:

1998‐2001. NOAA Technical Memorandum, NMFS‐SEFSC‐479, Miami, FL, 26 pp. 26 pp

NMFS (2006) Endangered and Threatened Species: Final Listing Determinations for

Elkhorn Coral and Staghorn Coral. Federal Register 71:26852‐26861

Patterson KL, Porter JW, Ritchie KE, Polson SW, Mueller E, Peters EC, Santavy DL, Smiths

GW (2002) The etiology of white pox, a lethal disease of the Caribbean elkhorn coral,

Acropora palmata. Proceedings Of The National Academy Of Sciences USA 99:8725‐8730

Peters EC, Halas JC, McCarty HB (1986) Calicoblastic neoplasms in Acropora palmata , with a

review of reports on anomalies of growth and form in corals. Journal of the National

Cancer Institute 76:895‐912

Porter JW, Dustan P, Jaap WC, Patterson KL, Kosmynin V, Meier OW, Patterson ME,

Parsons M (2001) Patterns of spread of coral disease in the Florida Keys. Hydrobiologia

460:1‐24

Robertson DR, Hoffman SG, Sheldon JM (1981) Availability of Space for the Territorial

Caribbean Damselfish Eupomacentrus planifrons. Ecology 62:1162‐1169

Rodriguez‐Martinez RE, Banaszak AT, Jordan‐Dahlgren E (2001) Necrotic patches affect

Acropora palmata (Scleractinia : acroporidae) in the Mexican Caribbean. Diseases Of

Aquatic Organisms 47:229‐234

Shinn EA (1989) Reefs of Florida and the Dry Tortugas : Miami to Key West, Florida, July 2‐

7, 1989. American Geophysical Union:45 pp.

33
Weil E, Hernández‐Delgado EA, Bruckner AW (2002) Distribution and Status of Acroporid

Coral (Scleractinia) Populations in Puerto Rico. In: Bruckner AW (ed) Proceedings of the

Caribbean Acropora Workshop potential application of the U.S. Endangered Species Act

as a conservation strategy: April 16‐18, 2002, Miami, Florida. NOAA Technical

Memorandum NMFS‐OPR‐24, Silver Spring, MD, pp 67‐94

Weil E, Losada F, Bone D (1984) Spatial variations in density and size of the echinoid

Diadema antillarum Philippi on some Venezuelan coral reefs. Bijdragen tot de Dierkunde

54:73‐82

Williams DE, Miller MW (2006) Importance of disease & predation to the growth &

survivorship of juvenile Acropora palmata & Acropora cervicornis: A demographic

approach. Proc 10th Int Coral Reef Symp:1096‐1104

34
GLOSSARY 

active (tissue loss) - describes disease lesions that are believed to be actively progressing 

based on exposed skeleton with no algal colonization and frequently irregular jagged 

lesion margins. 

attachment point - tissue & skeletal growth formed when a fragment fuses with the 

substrate. Fragments attaching to the substrate frequently produce several attachment 

points. 

base - portion of the colony encrusting the substrate 

bleached - Coral tissue that is lacking zooxanthellae- Tissue appears translucent but is 

intact. 

calcareous coralline algae (CCA) - light pink to burgundy encrusting algae with a calcified 

thallus also known as ‘pink pavement’. 

chimney - Growths of Acropora spp. tissue resulting from threespot damselfish bites. 

Repeated bites during tissue healing result in upward growth of tissue and skeleton 

which are frequently toped with filamentous algae. 

colony - Operationally defined as an area of continuous tissue OR disconnected patches of 

tissue found on the same underlying skeletal structure (presumed to be formerly 

continuous). 

corallite - cup shaped calcium carbonate structure secreted by and containing a single polyp 

filamentous algae -– Fast-growing algal forms that quickly colonize ‘new’ reef surfaces 

such as newly dead coral skeleton. The intensity of filamentous algal growth can be 

used to gauge the approximate age or rate of progression of an area of recent mortality 

fragment - broken portions of standing colonies found loose on the surrounding substrate. 

Once the fragment attaches to the substrate it can be considered (here) an ‘asexual 

recruit’. 

genet - the genetic individual resulting from a single planula (zygote). For clonal organisms 

such as Acropora a genet often includes numerous ramets (asexually produced 

individuals). Determining if colonies belong to the same genet requires genotyping in the 

lab and can only be guessed based on observations in the field. 

healing – re-growth of healthy tissue over a former lesion or scar. 

isolate - physiologically continuous patch of tissue regardless of size or shape. 

lesion - Area of recently exposed skeleton that is presumed to result from disease 

35
margin - boundary between live tissue and adjacent substrate (including Acropora

skeleton/dead areas) 

polyp - single unit of the coral colony contained within a corallite. 

ramet - products of asexual reproduction by a given ramet genet. 

recent (tissue loss) - Exposed Acropora skeleton with little or no colonization by 

filamentous algae and relatively sharp, observable corallite structure. 

recruit - smaller colonies (<20cm) are frequently described in the literature as ‘recruits’ 

since they are presumed to have appeared recently. In this study a recruit is a colony that 

as appears in the study plot only after the initial survey. It includes both fragments that 

have re-attached to the substrate (asexual recruits) and new colonies that appear from 

settled planulae (sexual recruits). 

remnant - describes patch(es) of tissue (isolates) that result from a larger colony that lost 

substantial tissue leaving only 1 or a few disjointed patches of tissue. Typically describes 

patches that have grown new branches since they were isolated. 

snails - In the context of this protocol, the term ‘snails’ refers only to those found to prey on 

Acropora, chiefly Coralliophila abbreviata but also Coralliophila caribbaea where 

present. 

36
Figure 1. a) While dense monospecific patches or “thickets” are characteristic of 

Acropora palmata, these complex structures are not conducive to the ‘demographic’

monitoring methods described here. ‘Demographic’ monitoring tracks the 

performance of individual colonies (b-c) over time. If colonies are densely spaced in a 

small area it becomes difficult to discern individuals from their neighboring colonies. 

There are distinct boundaries separating individual ‘colonies’ (b-c).

a)

b) c)
Figure 2. Schematic illustration of 

plot placement. a) an example 

based on a spur and groove type 

reef structure and b) from a 

fringing type reef. Acropora 

palmata colonies are represented 

by (∆) and (▲). Shaded triangles 

represent those that would be 

selected for tagging using 

randomly generated compass 

bearings and distances from the 

central plot stake (●). There are a 

total 35 colonies selected within 

the three plots. Tagged colonies 

in the plots on the far left and far 

right in both frames were selected 

randomly. All colonies in the 

center plot of both frames were 

tagged because there are less than 

12 colonies within the plot area. 

a)

b)
Figure 3. Remnant patches of live tissue on large colony skeletons represent 

challenges in designating what is an individual colony for monitoring. This 

designation is crucial to tracking changes in the colonies and recruitment so it 

is important to be consistent over time. Guidelines are presented in the text 

to help the surveyor make consistent decisions on the boundaries of a colony. 
Figure 4. Illustration of three rules used to distinguish colonies. In (a) live tissue is clearly 

continuous. In (b) live tissue is not physically connected, but the two patches of live tissue are 

on the same remnant structure, so the two patches should be considered the same colony. In 

(c), the three regions of live tissue should be considered separate colonies. Some ambiguity 

exists in distinguishing whether the remnant/dead structures of the remaining tissue patches are 

related, and because the tissue patch on the right is more than 1 meter from the larger tissue 

patch on the left, these two patches should be considered separate colonies. The small patch of 

live tissue on the lower left is within one meter of the larger live patch on the left, but is clearly 

not on the same remnant/dead structure, so should therefore be considered a separate colony. 

Fig D Colony structure
Figure 5. Field sheet used by Diver B to select the colonies to be tagged. On the right are sets of ran

coordinates that are followed by Diver B. If the center of a colony is found within 0.5m of this point Then the actual distance and degrees of the center of the tagged colony should be recorded with the left side of the field sheet. 
Fig E Fig F Colony 

Tagging

Figure 6. Tags should be nailed to an 

adjacent area of bare substrate, close 

enough to clearly identify the tagged 

colony, but far enough to allow for 

the tissue around the base to grow. 

Arrow indicates tag. Tags shown in 

frames a and b are appropriately 

spaced. The tag in frame c is too 

close to the base of the colony, 18 

months later the tag is being 

overgrown by the colony and adjacent 

Palythoa caribaeorum .

Figure F

a)

b)

c) d)

4/2004 10/2005
Figure 7. Field Sheet used by Diver A to record the initial survey of the colonies to be tagged. The fiein detail in the text. a) Fields have been filled in for 4 colonies at a hypothetical site called Horsesho

that the codes for several of the fields are listed at the bottom of the field sheet. 

a)

b)
Figure 8. Orientation of colony measurements of A) the maximum length and width, and B) 

height (perpendicular to the substrate) of a tagged colony.

FigG Fig H Colony Assess
L

W

H

Figure 9. Photos taken of a colony that was assessed as 100% live. All 4 sides are shown to emphthe colony that were included in the LxWxH measurement must be considered when estimating d show the orientation of the length (L), width (W) and Height (H) measurements. 

a) b)

c) d)
Figure 10. Two colonies shown from various angles to illustrate an example of a colony with an 

estimated a) 60% coverage of live tissue and b) 30% cover of live tissue. The yellow rulers are 

superimposed to demonstrate how the dimensions (length (L), Width (W) and height (H) were 

measured when estimating the %live for that colony. The meter stick (white pole) is used to take 

measurements in the field and to provide scale in plan view (first frame) images. Note that in 10-b 

the plan view photo is not directly over the colony making it hard to indicate the length and width 

of this colony. Furthermore, because two thirds of the colony is dead it is difficult to differentiate 

the colony margins. The meter stick was placed at the base of the colony to highlight the colony 

boundary. Note also that Height is measured from the lowest live point (at arrow in bottom frame) 

rather than extending to the seafloor. Both ways are valid as long as the % live takes this into 

account. 

a) 60% live b) 30% live

H

L

W

H

L

W
Figure 11. Healthy Acropora colonies are 

golden to orange brown in color (a). The tissue 

itself is translucent but harbors zooxanthellae 

which give it color. The calcium carbonate 

skeleton is bright white beneath the tissue (b). 

Colonies are comprised of many individual 

polyps which each secrete a cup shaped 

corallite. Polyps are withdrawn into the 

corallites during the day but often the 

tentacles protrude slightly above the corallite 

and can be seen on close inspection. Colony 

margins, especially branch tips are normally 

lighter in color where new tissue has grown 

but has not yet acquired zooxanthellae (c).

Live tissue

Bare skeleton

Corallites

Lighter color 

on growing 

margins

1.0 cm 10 cm

1.0 cm

a)

b) c)
L

L

R

R

R

R

R

Figure 12. When tissue is 

lost the bright white coral 

skeleton is exposed and 

gradually undergoes 

succession. In the first few 

weeks it is possible to 

roughly distinguish 

‘recently dead’ areas from 

‘long dead’ areas of a 

colony based on the 

relative amounts of algal 

colonization and skeletal 

erosion. ‘Recently dead’

(R) areas have light or no 

algal colonization and the 

corallite structures are 

still sharply defined. 

‘Long dead’ (L) areas have 

eroded corallites and 

heavier colonization by 

filamentous algae and/or 

fleshy or calcareous algae. 

In general the longer the 

skeleton has been exposed 

the darker it appears but 

it is helpful to look at the 

corallites (or lack of) to 

help distinguish recent and 

long dead.
a)

Figure 13. Successional 

changes from ‘recent 

dead’ (a) to ‘long dead’

on a single colony 

branch (b-d in order). 

Recent (a) and long 

dead (b) classification is 

qualitative and can not 

be precisely delineated. 

Observers should try to 

be consistent in their 

classification and take 

adequate photos that 

will permit verification 

later. 

b)

c) d)
Figure 14. Two examples of colonies with a recent mortality 

rank= 1. Relative to the size of these colonies the recent 

mortality is “present but very sparse” (see text). 
Figure 15. Two examples of colonies with a recent mortality rank= 2. Relative to 

the size of these colonies the recent mortality is “moderate(10-25%)” (see text). 
Figure 16. Examples of colonies with a 

recent mortality rank= 3. Relative to the 

size of these colonies the recent 

mortality is “High, affecting 30-55% of 

the colony live area ” (see text). 

d)
Figure 17. Examples of colonies with a recent 

mortality rank= 4. Relative to the size of these 

colonies the recent mortality is “Catastrophic 

affecting, 60-85% of the colony’s live area” (see 

text).
Figure 18. Examples of colonies with a recent 

mortality rank= 5. Relative to the size of these 

colonies the recent mortality affects almost 

the “entire colony >/= 85%” of the colony’s live 

area” (see text).Fortunately this degree of 

recent mortality is rarely observed. 
Live Tissue

Active 

Tissue 

Loss

Recent Tissue Loss

Figure 19. Exposed skeleton that is bright white with NO algal colonization frequently has a 

jagged margin (a) indicating that tissue loss is actively progressing. The irregular margin 

results from small pieces of tissue that are still clinging to the skeleton (only seen on close 

inspection) (b). For the purposes of the colony condition assessment described in this 

protocol ‘active’ tissue loss is not distinguished from ‘recent’ tissue loss, but recognizing it 

can help recognize the pattern and rate of tissue loss which aids in assigning it to the 

appropriate type. a) band-like pattern of tissue loss is relatively uniform; c) patchy tissue 

loss is exposed skeleton typically surrounded by live tissue. 

1.0 cm

a)

Active 

Tissue 

Loss

10 cm

d)

Note small scattered remnants of live tissue 

clinging to recently exposed skeleton

1.0 cm b)

10 cm

c)
a)

b)

Figure 20. White band disease has a distinctive band of recent mortality. 

The bright white band of exposed skeleton grades into increasingly algal 

colonized skeleton indicating that the band is progressing gradually along 

the branch. Typically the band progresses from the base towards the tips (a) 

and is often progressing evenly up adjacent branches (a-b) 
Figure 21. In the Florida Keys, WBD is often observed to advance from the base of the colony along th

branches. The disease front often stalls as it reaches the branch margins, leaving branches with a ‘whviewed from above (Reproduced from Williams & Miller 2006). A) and B) show time series of two sepa
Figure 22. White pox type 

recent mortality is 

characterized by multifocal

and irregular patches of 

exposed skeleton or ‘lesions’

with rates of tissue loss of 12 

cm2 per day. Lesions’ can be 

found on any part of the 

colony 
Figure 23. a) recent mortality classified as white Pox (WPx) lesions. b) lesions are healing, 

note the white lip around the advancing tissue which is typical of new tissue growth. In 

frames b-d the original lesions are observed to heal completely. c) two additional lesions 

in the bottom right have appeared since the last photo but are also in the process of 

healing which continues in d; small white spots termed damselfish chimneys also appear, 

the threespot damselfish is also visible in frame c. 

a)

d)

b)

c)
Figure 24. Rapid Tissue Loss 

(RTL) is a general descriptive 

term used to avoid the use of 

specific disease ‘diagnoses’ for 

cases such as illustrated here 

that have large irregular lesions 

that do not conform to the 

characteristic patterns of either 

White Band or WPx types of 

lesions. 
a) b)

c) e)

d)

Figure 25. Manifestations of rapid 

tissue loss (RTL) vary greatly from poxlike multi-focal lesions (a) to band-like 

progression (b), however in all cases 

RTL progresses more rapidly than has 

been documented for WBD or WPx. In 

most cases the pattern of recent 

mortality is highly irregular (c-e).
Figure 26. Coralliophila 

abbreviata feeds on Acropora

spp. (and other coral) tissue 

leaving an area of bare 

skeleton (recent mortality) 

called a feeding scar (FS). 

They are often found feeding 

in a group (typically 3-6 snails) 

but algal colonization on their 

shell makes the highly cryptic, 

resting on dead substrate at 

the base of colonies or 

undersides of branches when 

not actively feeding. They 

usually remain on a single 

colony for extended periods of 

time (months or longer). While 

they are often found feeding 

(b) but when not feeding they 

migrate away from the live 

tissue and are found ‘resting’

(c) and outside of the feeding 

scar. Note the scalloped 

pattern at the margin of the 

feeding scar and live tissue is 

apparent in frames a and c. 

a)

b)

a)

b) c)

FS

FS

FS
Figure 27. Coralliophila abbreviata

predation typically leaves a path or 

trail of denuded skeleton behind. a) 

shows a trail of recent mortality cased 

by snail feeding (i.e. Feeding scar 

(FS)), and older dead areas on the 

branch which may be areas fed on by 

snails in the past. These snails appear 

to have encountered the branch tip 

and turned back a few times in the 

area shown in this frame. However, 

only the bright white area (outlined in 

blue) would be considered recent 

mortality and assigned FS as type, the 

rest is speculation! b) extensive ‘snail 

trails’ can be seen on larger colonies, 

the dotted yellow line shows a 

probable trail created by snails (at 

arrows) feeding over the past 6-8 

months. 

a)

b)
a)

b)

c)

Figure 28. Coralliophila abbreviata

are only rarely found feeding in an 

area entirely surrounded by live 

tissue (a-b). In general, snails 

should be excluded as the cause of 

lesions completely surrounded by 

live tissue (e.g. white pox) unless 

the snail is observed. c) snails (at 

arrow) commonly feed on the 

underside of a branch resulting in a 

band-like FS that might easily be 

mistaken for WBD. (see text for 

further details)
Figure 29. Parrotfish typically feed 

by scraping reef substrate for the 

endolithic algae and sponges. 

However scattered bite marks (a) 

are not uncommon on live A. 

palmata. Scrapes are more common 

on raised parts of the colony and 

branch tips (b). Occasionally the 

biting becomes more intense and 

concentrated in a behavior known as 

‘focused biting’ (c). 

a)

b)

c)
Figure 30. a) Focused biting by Parrotfish (Sparisoma viride) can result in 

substantial areas of tissue loss and skeletal damage. These bites are usually 

focused on ridges, sides of branches or other protruding portions of a colony (at 

arrows). b) However colonies are able to heal rapidly, the pictured colony healed 

completely in less than 6 months (in fact a new proto branch has formed at the 

upper left arrow). 

a) Figure Z

b)

11/21/2002

5/12/2003
Figure 31. Colonies are 

occasionally found with the 

characteristic appearance of 

focused biting (b), however 

on closer inspection the 

white marks have intact 

corallites and bleached 

polyps (a). Since the tissue is 

intact, it should not be 

considered as recent 

mortality, instead it will be 

recorded as a type of 

bleaching. 

a)

b)
Figure 32. a) A dark greenish black band of ciliates is sometimes seen associate with recent 

mortality. (b) The ciliates vary in density, forming a thick fuzz resembling black band 

disease in some cases c) The band appears more sparse in areas of faster tissue retreat.

b)

a)

c)
Figure 33. Cilliate bands are very 

conspicuous but may co-occur 

with less distinctive types of 

recent mortality. A) when a 

cilliate band occurs with a white 

band of recent mortality it is 

impossible to know if the cilliates

are the sole cause of the 

mortality or if white band disease 

is the primary cause and cilliates

are a secondary ‘cause’. b) Snail 

feeding scar is the primary type of 

recent mortality and c) cilliates

are secondary.

b)

a)

c)
b)

a)

c)

Figure 34. a) Cliona sp.? Is very 

common on reef substrates 

including long dead A. palmata

skeleton and is visible only 

where its ostia protrude (at 

arrows). b) Ostia can also 

protrude through live tissue 

resulting in a lesion. c) this 

fresh branch break shows a 

cross section through an 

excavated branch. The ostia

can be seen (at arrows) along 

with the main body of the 

sponge. 
b)

a)

c)

Figure 35. a) When fish fecal material (Fish 

Poop) lands on A. palmata’s flat branches 

and remains there, the tissue underneath 

can get smothered resulting in b) scattered 

irregular lesions that can resemble a white 

pox lesions. c) Fish poop lesions are often 

crescent shaped and may have traces of 

fish poop among the denuded corallites. 
Figure 36. Bleaching occurs when 

zooxanthellae are lost from 

intact coral tissue (a). Bleached 

tissue is still alive and appears 

translucent on close inspection 

(b). Note that polyps are visible 

both in the normal and bleached 

tissue (b, at arrow) indicating 

that the tissue is still alive in this 

portion of the colony.

b)

a)
b)

a) c)

Figure 37. Bleaching may 

affect whole colonies but 

frequently affects only 

parts of a colony. In this 

protocol bleaching includes 

colonies with partial 

bleaching (a-b) and 

uniformly pale colonies (c). 
Figure 38. Wedge-shaped area 

of bleached tissue (known 

informally as ‘Wedgies’) are 

usually found around the base 

of a colony leading to the 

tissue margin (a-b), but they 

can occasionally be found in in 

the interior of the colony 

surrounded by live tissue (c). 

These marks persist nearly 

unchanged for at least several 

years. 

a)

b) c)
Figure 39. Ring-shaped bleached areas 

are thought to be caused by yellowtail 

damselfish bites. These marks routinely 

appear and heal on a colony b) After 2 

weeks some of the rings present in 

frame b are still present while others 

have disappeared and new rings have 

appeared. 

a)

b) 6/25

c) 7/10
Figure 40. Threespot Damselfish maintain 

territories on A. palmata (b). They repeatedly 

bite the tissue, clearing space for algal growth. 

As tissue begins to heal over the bite, it is 

repeatedly interrupted by more bites. The result 

is a ‘chimney’ (a) which is a small tube like 

protuberance, often open at the top and capped 

with algae. The tissue at the top of the chimney 

is often white like other areas of new Acropora

growth. The freshly bitten areas are also white 

but are flush with the surrounding tissue. 

a)

b)
Figure 41. Time series of a threespot damselfish territory over a two year period. A few 

chimneys are present around the margins, and though they change in number at each 

survey, the overall impact on the Acropora colony remains fairly constant. The chimneys 

are surrounding the damselfish’s ‘lawn’ (L) which is an algal turf covered part of the 

colony. While we can’t assume that the fish was the original cause of the tissue loss on 

this colony, it is preventing regrowth of tissue in that area thus the whole area, lawn + 

chimneys should be included when ranking the impact on this colony. Note that this 

territory is located at the base of a colony which is somewhat unusual).

4/04

7/04

10/04

1/05

4/05

8/05

11/05

2/06

L
Figure 42. Male threespot damselfish guard eggs 

laid within their territory. Nests are typically on 

the underside of a branch and surrounded by live 

Acropora tissue (a-b). A given nest seems to be 

used repeatedly, frame c and d were taken more 

than a year apart. When eggs are present the 

nest will have an grey iridescent cast, often over 

a very fine deep red filamentous algae turf. The 

area occupied by a nest should be included in 

the damselfish impact rank, though it is usually 

minor compared to the chimney and lawn 

impacts. 

‘Empty‘ nest Nest with eggs

a)

c) d)

b)
Figure 43. Examples of damselfish impact with rank=1 

(a-b) and rank=2 (c).

a)

c)

b)
a)

Figure 44. Examples of damselfish impact ranks 3 (a-b) and 4 (c). 

b)
Figure 45. Examples of damselfish impact rank 5.

a)

b)
Figure 46. Irregular growth that is 

relatively smooth and depleted in 

zooxanthellae is termed neoplastic

growth (a-b) and recorded as ‘NP’ in 

the skeletal anomaly column of the 

field sheet. Irregular growth that has 

zooxanthellae and defined, albeit 

abnormal, corallites is termed 

hyperplastic growth and recorded as 

‘HP’ (c-e). 

a) b)

c) d)

e)
Figure 47. Colonies are 

occasionally observed with a band 

of white, densely packed of 

corallites. There is no recent 

mortality associated with the 

bands however it gradually 

retreats across the colony. These 

bands are closely associated with 

yellowtail damselfish (c), and 

may be the result of repeated 

nesting activity. 

a)

b)

c)
a)

Overgrowth by 

brown clionid 

sponge

b)

Figure 48. Cliona langae is a very thin encrusting sponge that aggressively 

competes with or invades coral tissue and can readily overgrow it (a-b). 

It has a velvet feel and is chocolate to olive brown in color. Note that the 

white tissue at the margin is not recent mortality. It is more accurately 

characterized as a tissue rejection response. 
a)

Overgrowth by 

brown clionid 

sponge

Figure 49. Erythropodium caribaeorum is 

an encrusting tan colored gorgonian (a) 

that overgrows Acropora tissue. When its 

polyps are extended its long tentacles give 

it a ‘furry’ (b) look (note that the dark 

greenish-black area is the result of cilliate

infestation and not part of the gorgonian). 

When retracted it has a leathery 

appearance (c). Acropora is an excellent 

competitor and often overgrows other 

sessile organisms (d). 

a) b)

c)

d)
Figure 50. Fragmentation is an important aspect of Acropora life history, allowing rapid clonal 

reproduction. Fragmentation is scored as the number and ranked size (diameter) of branches 

that have broken off, as determined by examination of the broken scars on the focal colony. 

In the top picture, the scars have a greenish cast due to colonization of algal turf following an 

acute disturbance event.

a)

b) c)
www.jaxshells.org www.jaxshells.org

www.jaxshells.org www.jaxshells.org

T. deltoidea C. abbreviata

Figure 51b. Coralliophila abbreviata: Feeds 

on coral tissue typically found in a small 

group associated with live tissue or a 

feeding scar. Shell has fine ridges that are 

sometimes visible through the encrusting 

algae. Bright yellow body and gold 

operculum. C. caribbaea (shown in frame 

a) is smaller at full size and has a white or 

tan body with a burgundy operculum. They 

are also found in a small group and near a 

feeding scar.

Figure 51. Coralliophila 

abbreviata and C 

caribbaea are corallivores

commonly found feeding or 

resting on Acropora

colonies. It is important to 

distinguish them from other 

common snails (e.g. T. 

deltoidea) that do not feed 

on coral so that an 

accurate census of these 

predators can be done. C. 

abbreviata is more common 

on Acropora than C. 

caribbaea. These three 

species can be 

distinguished based on 

some physical traits and 

habits. 

Figure 51c. Thais deltoidea: Feeds on other 

snails and is commonly found on reef 

substrate including the dead portions of 

Acropora colonies. Only coincidentally 

associated with live coral tissue. Dull knobs 

on the outer whorl that are subtly visible 

through the encrusting algae. Burgundy body 

and very dark purple/brown operculum. 

C. caribbaea

a)

b)

c)
1

2

3 Figure 52. Threespot damselfish (Stegastes planifrons) adults are 

similar in appearance to Dusky, 

Beaugregory & Cocoa damselfishes. 

Their body color varies from light to 

dark yellowish brown. Their most 

reliable distinguishing features are 

the spots & behavior: a) They have 

three spots: 1. bright yellow eyelid; 

2. black & yellow patch at base of 

pectoral fin; 3. black patch at base 

of caudal fin. At least one of these 

will be visible (note that the cocoa 

has a similar spot on the base of the 

caudal fin). c) They are very 

territorial and often will aggressively 

approach observers, particularly 

around egg masses. Other species of 

damselfish tend to be less aggressive 

but this varies within a species. c) 

Juveniles are bright yellow with 2 

black spots & not aggressive. 

a)

c)

b)
Figure 53. Yellowtail Damselfish (Microspathodon chrysurus) are 

easily distinguished by their 

yellow tail as adults (a) and 

vivid blue spots as juveniles (b). 

They are often found sheltered 

among the branches (c). While 

they are closely associated with 

A. palmata, they often ‘patrol’

a territory that may include 

more than one colony. Frame a 

also shows a nest, note that it 

is surrounded by a white band 

(see also Figure 47) that should 

not be confused with white 

band disease (see Figure 20).

b)

a)

c)
Figure 54. a) Schematic illustration of the plot mapping procedure. b) Diver A records colony information and c) Diver B reads the bearing to each colony designated by Diver A.

a)

b)

c)
Figure 55. Data collection is divided between a) diver A and b) Diver B for the quarterly 

surveys. 

a)

b)
APPENDICES 
1° 2° 3° 1° 2° Ad Juv Ad Juv sm med lg xlg

Overgrowth

Additional Notes:

H

% Live

* Recent mortality types: WPx= white pox, WBD= white-band, RTL= rapid tissue loss, FS=snail feeding scar, CB=cilliate band, PFB= parr

FP=fish poop, Unkn= unknown ** Bleaching types: Bl= bleached, R= rings, W= wedgies ***Skeletal anomolies: Hp=hyperplasia, Np=n

Unkn=Unknown

Diver A: Initial Colony SurveySite________________________ Plot _______ Date____________________

Recent 

mort*

RankTypes

X

Tag # L W

Bleaching**

 # Snails

# 3-Spot 

damsels

# Yellowtail 

damsels

RankTypes

Damselfish 

Impact (Rank)

Skel anoms***

 # Branch 

Breaks
Random Coordinates:

Site_______________

Stake Depth: Deg Dist Deg Dist

Stake Waypoint# 75 3.0 300 4.5

Plot Colony Tag # Deg Dist Notes (tag loc., depth, etc) 135 4.5 35 3.0

1 1 260 5.0 105 3.5

1 2 135 1.5 215 0.5

1 3 110 3.5 240 4.0

1 4 190 0.5 30 4.5

1 5 275 1.5 10 0.5

1 6 175 4.0 85 2.0

1 7 140 2.5 125 7.0

1 8 200 6.5 225 1.5

1 9 225 5.0 310 2.5

1 10 225 7.0 30 1.0

1 11 270 4.5 280 3.5

1 12 40 5.5 125 6.5

280 1.5 245 6.5

345 1.0 150 5.0

60 3.0 15 5.0

30 1.5 20 5.0

Site_______________ 175 4.0 260 1.5

Stake Depth: 115 3.5 290 2.0

Stake Waypoint# 35 2.0 20 0.5

Plot Colony Tag # Deg Dist Notes (tag loc., depth, etc) 235 6.0 20 6.5

2 1 85 5.0 300 3.0

2 2 40 2.0 215 0.5

2 3 110 2.0 80 0.5

2 4 190 6.0 305 3.5

2 5 0 1.5 230 6.5

2 6 150 2.5 20 4.0

2 7 225 5.5 65 7.0

2 8 110 7.0 300 0.5

2 9 230 7.0 340 2.5

2 10 55 4.5 50 5.5

2 11 205 1.5 30 7.0

2 12 85 2.5 215 3.5

185 2.5 275 2.5

335 5.0 195 1.5

295 2.0 20 2.0

Notes: 285 1.0 105 6.5

165 2.0 350 7.0

195 7.0 50 6.5

155 4.0 80 4.0

205 4.5 5 1.0

150 3.5 155 3.0

320 6.0 170 4.5

325 4.5 295 1.0

295 1.5 135 3.0

220 5.5 55 4.0

Diver B Tagged Colony Selection

Date________________

Date________________ Plot 1 Plot 2

A-2
Date____________________

1° 2° 3° 1° 2°

Date____________________

1° 2° 3° 1° 2°

Diver A Quarterly Colony Condition Assessment

Site________________________ Plot _______

X

Tag #

LWH

% Live

Recent 

mort*

Bleaching**

Skel anoms***

Notes

Rank

Types

Rank

Types

Damselfish 

Impact (Rank)

Site________________________ Plot _______

H

% Live

Recent 

mort*

Bleaching*

*

X

Tag #

L W

Skel anoms***

Notes

Rank

Types

Rank

Types

Damselfish 

Impact (Rank)

* Recent mortality types: WPx= white pox, WBD= white-band, RTL= rapid tissue loss, FS=snail feeding scar, CB=cilliate band, 

PFB= parrrotfish bites, CL=Cliona lesion, FP=fish poop, Unkn= unknown ** Bleaching types: Bl= bleached, R= rings, W= 

wedgies ***Skeletal anomolies: Hp=hyperplasia, Np=neoplasia, DWP=dense white polyps, Unkn=Unknown A-3
Site_________________ Plot_______ Date____________

Ad Juv Ad Juv sm med lg xlg

Site_______________ Plot_______ Date__________

Ad Juv Ad Juv sm med lg xlg

X

Tag #DegreeDistance

# Snails

# 3-Spot damsels

Notes

# Snails

# 3-Spot damsels

Notes

X

Tag #DegreeDistance

Diver B Quarterly Snail and Fish Census# Ylwtail damsels

 # Branch 

Breaks

# Ylwtail damsels

 # Branch 

Breaks

A-4
Annual Plot Map (Diver A: Distance from stake, colony description)

Plot_________________ Date____________________

Untagged 

Col # Deg Dist L W H % Live Colony Type** Untagged Col # Deg Dist L W H % Live Colony Type** 

X1 X17X2 X18X3 X19X4 X20X5 X21X6 X22X7 X23X8 X24X9 X25

X10 X26X11 X27X12 X28X13 X29 X14 X30X15 X31X16 X32

Plot_________________ Date____________________

Untagged 

Col # Deg Dist L W H % Live Colony Type** Untagged Col # Deg Dist L W H % Live Colony Type** 

X1 X17X2 X18X3 X19X4 X20X5 X21X6 X22X7 X23X8 X24X9 X25

X10 X26X11 X27X12 X28X13 X29 X14 X30X15 X31X16 X32

* Transcribe from Diver 2's data sheet after the dive

**BC= Branched Colony, RC=Remnant Crust, AF= Attached Fragment, SF=Stable Fragment LF=Loose Fra
Annual Plot Map (Diver B: Compass Heading)

Plot_________________ Date____________________ Plot_________________ Date__________

Untagged 

Col # Deg Untagged Col # Deg Tag # Deg Untagged Col # Deg Untagged Col # 

X1 X16 X1 X16X2 X17 X2 X17X3 X18 X3 X18X4 X19 X4 X19X5 X20 X5 X20X6 X21 X6 X21X7 X22 X7 X22X8 X23 X8 X23X9 X24 X9 X24

X10 X25 X10 X25X11 X26 X11 X26X12 X27 X12 X27X13 X28 X13 X28X14 X29 X14 X29X15 X30 X15 X30

Plot_________________ Date____________________ Plot_________________ Date__________

Untagged 

Col # Deg Untagged Col # Deg Tag # Deg Untagged Col # Deg Untagged Col # 

X1 X16 X1 X16X2 X17 X2 X17X3 X18 X3 X18X4 X19 X4 X19X5 X20 X5 X20X6 X21 X6 X21X7 X22 X7 X22X8 X23 X8 X23X9 X24 X9 X24

X10 X25 X10 X25X11 X26 X11 X26X12 X27 X12 X27X13 X28 X13 X28X14 X29 X14 X29X15 X30 X15 X30

Confirm previously measured coordinates:Confirm previously measured coordinates:
Annual Plot Map Plot Date
1

2

4

3

5

8

9

13

1. Metal stakes 

2. Sledge hammer

3. Aluminum numbered tags 

4. Cable ties 

5. Nails 

6. Transect line (10m length) 

7. Dive compass

8. GPS with waterproof case

9. PVC meter stick (marked 5cm)

10. Pencils

11. Rubber bands/string 

12. Aluminum clipboards

13. Dura Copy underwater paper

Key

6

10-12

7

B-1
Photoboard: for scale and labeling in field ph

Metriclose itself wide 

Front 

12” 

31/4” 

PVC Bocolor re

Bold, individual letters & numbers on front indicate colony or sample number in photo 

Small distributed fishing weights eliminate buoyancy and provide balance & stability 

Letters & numbers not being used are stored on the back the board 
Materials: 

1) Metric Ruler (plastic) 2) PVC sheet (0.3mm thick) 

3) PVC board (2.5mm thick) 4) Adhesive Velcro (Heavy 

Duty) 

 (two 12”x 2” strips) 5) Fishing Weights (6 x 1.5oz) 6) Super Glue 

7) Box cutter 8) Scissors 9) Cable ties 

10) Sharpie (Wide/Chisel Tip) 11) Drill (not shown) 

Instructions: 

A. Cut 2.5mm thick PVC to 12” x 3 1⁄4” using box cutter (score deeply then b

B. Glue metric ruler flush with the top edge of the board. C. Affix one strip of Velcro to board below the ruler D.Affix second strip of Velcro to the back side of the board (centered) 

IMPORTANT: Both sides of the board must have the same side (hook 

of velcro, the mate to both strips will be used later. 

E. Drill holes 2 on each end and 2 along the bottom edge. Each pair shoul

F. Attach fishing weights to the board by lacing the cable ties through the 

G.Cut remaining two strips of Velcro in half lengthwise (1” x 12” strips) H.Affix Velcro strip just below (<1/8”) the top margin of the 0.3mm thick P

sheet just below the Velcro. Repeat this with the remaining 3 strips of V

I. Use the Sharpie to write the appropriate letters and numbers and cut to 

individual letters. 

J. Stick the letters not being used on the Velcro on the back side of the bo