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Index | Overview | Methodology | Coral standards | Fish standards | Calibration | Data processing | Data sheets
AGRRA
Methodology
version 4.0, June 2005
(Revision by Philip Kramer, Judith Lang, Kenneth Marks, Rodrigo
Garza-Perez & Robert Ginsburg)
Equipment
Selection of reefs and sites
Corals, algae and Diadema
Coral reef fishes
Optional components
The following equipment is required for each diver in addition to basic snorkeling gear and SCUBA gear (including depth gauge):
Note: If you normally use prescription
lenses to correct your vision, it is
very important to have a diving mask with the correct prescription lenses.
Otherwise you may not be able to distinguish some important details while doing
the surveys.
Stony corals,
Algae, and Diadema:
1. Underwater datasheets.
Attach
the datasheets onto a clipboard, underwater slate or writing cylinder (see below).
The Appendix has an example of the AGRRA benthos datasheet template. You can
photocopy the data template onto both sides of white underwater Duracopy paper
(boxes of 100 sheets, contact J.L. Darling Corporation, phone: (253) 922-5000,
fax: (253) 922-5300; address:
2. A 10-m long transect line.
A 10-m polypropylene line marked at 1 m intervals (with cables-ties, electrical tape or permanent ink) to which a small dive weight has been attached at each end. Or, whenever available, use a lead-core (Duraflex) weighted line. Please ask your local fishing supplies store/company; if they have it, they should know what it is.
3. A 1-m long measuring pole.
A 1-m long PVC pipe (~ ½ ” diameter) marked in 10-cm intervals.
4. A 25 x 25 cm quadrat (25 cm inner diameter)
Construct
quadrats by gluing together ¼’ or ½" PVC water pipe and elbows and drill
two holes on each side to let the air out.
5. A small plastic ruler tied to the clipboard, slate or writing cylinder or attached to your wrist with a series of interconnected rubber bands. Trim the ruler to have a narrow, tapered point, but still be legible, at the basal 5 cm.
For convenience, wrap the transect line tightly around the quadrat; insert the meter pole and datasheet down the center between the line. Please consult the Appendix for the pictures of the gear.
Fish
1. Underwater datasheets.
Attach the datasheet onto a clipboard, underwater slate or writing cylinder (see below). The Appendix has an example of the AGRRA fish datasheet template. Datasheets for REEF rover diver surveys are available for $0.60/page by ordering from REEF at (305) 451-0312.
2. Transect tape and weight.
A 30-m fiberglass transect tape with a 2-3 lb weight attached at one end of the line. Commercially available PVC surveying tapes are suitable for the transect line. A clip can be attached to the reel and suspended from the diver’s belt, which allows for the tape to deploy freely as the diver swims.
3. A graduated T-bar or other measuring device (for fish density counts).
Construct a T-bar using ½" diameter PVC pipe and a T connector (available at hardware stores). It has a 60 cm long handle and two equal-length arms providing a total width across the top of 1 m. Paint a scale along the arms showing 10-cm increments. The slate can be mounted on the T-bar to facilitate carrying the equipment. (See Apendix for details).
Writing cylinders
A writing cylinder is a "thick walled" (1/4" thick) PVC pipe that is 4" in internal diameter by ~18 cm long, with 3 holes drilled near one end through which surgical tubing is strung to fit over your wrist. The advantage is that it keeps hands free to hold other surveying equipment and to hold on in strong surge or waves. A datasheet is attached to the outside of the cylinder with tape.
selection
of reefs and sites
For the purposes of AGRRA, a REGION is defined as the coarsest scale category (~100-1000 km scale); followed by an AREA (~10-100 km scale); a REEF (~1-10 km scale); and a SITE (0.2 km scale). We recognize that reefs vary greatly in size, complexity, depth, profile, and coverage per km of coastline throughout the region. What follows are our recommended procedures for selecting survey sites, however we fully understand that it may be necessary to modify these procedures to accommodate the special conditions of each large-scale assessment. It is vital for the success of AGRRA that these procedures must be followed as closely as possible, and that all modifications to them be carefully noted when the data are compiled.
Reef
Selection
The method for selection of REEFS to assess will be influenced in part by their abundance and distribution in your area and the amount of sampling effort to be undertaken. For most AGRRA surveys, the goal is to characterize the REEFS within a particular AREA (e.g., the sampling domain). Therefore, it is suggested that the limits of the AREA be delineated using either pre-defined biogeographic divisions (seascapes) or natural geography (island, atoll, bank, etc.). If the extent and/or number of REEFS (e.g., fringing, patch, barrier) within your AREA is so limited that they can all be assessed in a reasonable time frame there is no problem. However, if the extent, number and/or habitat complexity of REEFS are large, then they should be subdivided or “stratified” and representative examples randomly selected from each subdivision (e.g., the apples, oranges and bananas approach).
The most obvious stratifiers are
geomorphic characteristics of reef habitats that are influenced by cross-shelf
position, orientation, depth, slope, etc. When choosing reef habitats to
survey, try to avoid hard-grounds, pavements and other habitats that lack a
framework constructed of reef-building corals. Two of the most ubiquitous and
important zones within typical fringing or bank-margin
For each REEF type that is chosen, you should try to survey one SITE within each chosen depth interval, whenever both habitats are present, even if most of the A, palmata in the reef crest are dead and/or their colony borders are unclear–in which case substitute point counts for individual-coral assessments as described below.
Site Selection
A SITE is defined as an area of habitat that is more or less homogeneous and accessible from a boat anchored or moored in one place. Spatially, a SITE is roughly a 200m x 200m square unit. Once reefs are stratified, the idea is to either select representative SITES based on local knowledge (strategic) or to select SITES randomly (unbiased). For the latter method, give each reef within a subdivision a number and use a random method to select the ones to assess. If there are no clear bases for making subdivisions (e.g., in a continuous bank-barrier or fringing reef several kilometers long), then SITES should be located using a grid superimposed over the sampling AREA following the generalized random tessellation sampling (GRTS) approach. Sampling units (generally hexagons or squares) can be generated using a variety of ESRI Arc-View GIS extensions. An ESRI extension called SPOT originally created for MARXAN conservation planning is well suited for this type of hexagon generation. The size and number of units should be adjusted in size to the number of SITES that will be surveyed (effort) for each reef type (stratum). Random SITES can be determined within each sampling unit using Arc-View extension Random Point Generator available from www.esri.com. It is suggested that 2 alternative SITES also be generated within each sampling unit in case the primary SITE is determined unsuitable (for example, the bottom type is misclassified or the SITE is too dangerous to survey).
Depending on the methods and resources available for your use, REEFS that are selected will generally fall into one of three categories:
1. Unbiased- chosen based on a random sampling strategy;
2. Strategic- chosen with local knowledge because they are threatened, suspected to be degraded, or in particularly good condition.
It is critical that
the exact location of the actual survey be recorded using a GPS. In cases where
the survey takes place immediately below an anchored boat, simply record the
position of the boat once its position has stabilized. If the survey will occur
some distance from the boat (typically the case when surveying a reef crest),
note the distance and direction from the recorded GPS position so that the
position can be corrected later.
corals,
algae and diadema
1. At each SITE, record the following information on your UW datasheet before each dive. (We strongly suggest that each team member fills in every category.)
· Name of recorder as a four letter code (use the first two letters each of your first and last names);
· Date as day with two digits/abbreviation of month/year with two digits;
· Site Name (= local site name);
· Day Number (if during an expedition);
·
Site Number for that day;
· Latitude;
· Longitude;
· AGRRA location code as a three letter + three digit code (Use three letters from the name of the reef/expedition and number consecutively from 001 up to as many sites as are evaluated at that reef/during that expedition, e.g., sites at Little Island could be called LIS001, LIS002, LIS003, and so on);
·
Reef Type (e.g., fringing, bank, atoll, patch);
· How selected? (e.g., stratified random, stratified strategic);
· Reef Zone/Habitat (e.g., reef crest, reef front, spur and groove, bommies, platform, etc.).
2. In Time Start, record the time at which you start the first transect.
Haphazardly
lay the 10-m transect line just above the reef surface. Make sure the line is
taut. (Remember to complete the Site Description as described above).
Note:
Be sure to avoid and don’t cross any other transects that are being set
by your companions. Stay away from the edges of the reef. Also try to avoid
areas with abrupt changes in slope, deep grooves, large patches of sand or unconsolidated
coral rubble. You want to place the transect in areas where corals are likely
to grow, but once you are in an appropriate spot, don’t bias your selection:
instead swim without looking down at the bottom as you unreel the line. Unusual
reef features should only be included to the extent appropriate to their
relative abundance at the site.
The benthos survey can be made in three “passes” of the transect line as follows:
First pass:
3. Using the 1-m measuring pole for scale, swim a belt transect along the 10-m line. As you go, straighten any conspicuous kinks in the transect line. Count every “juvenile” and “adult” Diadema (juvenile and adult) that you can see within 1/2 m of each side of the transect line. Consider as juvenile all those Diadema that still show the black and white banded pattern on the spines. (Ignore all other species of sea urchins.) Count also any spiny lobster (Panulirus argus) and queen conch (Strombus gigas) and note their numbers in General Comments. (For the latter you may need to temporarily turn over shells that are upside-down to look for a living animal.) Because Diadema and Panulirus are cryptic, you must inspect all shelter-providing spaces along the line, so be prepared to poke your head under the bases of large corals or into crevices.
Second pass:
4. As you swim from one end of the transect line to the other, assess the cover, size and condition of each stony coral that is 10 cm in length or greater and for which any live or dead part of its skeleton underlies the transect line:
a. Identify the species (scleractinians, Millepora complanata, M. squarrosa, colonies of M. alcicornis that are encrusting other stony corals or the substratum + bases only of colonies that are encrusting gorgonians). Include all entirely dead colonies that can be identified at least to genus.
b. Measure the live tissue
cover of the colony under the line (cm) as if seen in a photograph, from above
in planar view. Record to the nearest cm for smaller corals (up to 50-100 cm) and to the nearest 5 cm for larger
(>100 cm) corals.
c. Measure the x, y, z
dimensions of the colony with the 1m pole: i.e.
the maximum length (x) and the maximum width (y) of the outward-facing
colony surface (both perpendicular to the axis of growth) as seen from above in
planar view, and the maximum height (z)
(parallel to the axis of growth) as seen from the side of the colony. Record
these measurements to the nearest cm for smaller corals (up to 25 cm) and to the nearest 5 cm for larger
corals. (Exact size measurements are less critical here since the colonies are
later grouped into size classes for comparative analyses).
Note: Colony boundaries can be difficult to
recognize when parts of the coral have died and are overgrown by other
organisms–particularly other colonies of the same species. Look for connected
live tissues, connected skeletal deposits above a common base, and at the size
and color of separated polyps.
Colonies derived from new recruits:
1) Live tissue, generally concentric with clear
edge boundaries. Often have a raised “lip” at edges approximately 1-2 mm above
underlying substrate/old dead coral.
2) Upward growth, branching evident.
3) Underlying substrate is very old dead.
Colonies derived from resheeting:
1) Live, often with preferred growth in one
direction, edges on at least one side often “merge” with underlying
substrate/dead coral.
2) Live tissue rarely displays upwards growth
(branching) except at tips.
d. Estimate the partial mortality (old and recent) of the colony surface from a planar view perpendicular to the axis of growth. Try to round your percentage to the nearest 5% unless it is very small or very large, in which case try to round to the nearest whole number (e.g., 1%, 97%). Although most colonies have some dead areas, 0% is recorded whenever these are restricted to the sides or bases and not visible when the outward-facing colony surface is viewed from above.
"Old dead" is defined as any non-living parts of the coral in which the corallite structures are either gone or covered over by organisms that are not easily removed (certain algae and invertebrates). If it is entirely “old dead”, indicate this on your data sheet as 100% “old death”, as long as you can identify it to either to the species (e.g., Acropora palmata by gross morphology; Montastraea cavernosa by polyp size and shape) or to the genus (e.g., Diploria by size of meandering ridges and valleys).
Note: In some cases, a coral may
be partially or completely overgrown by one of the species of brown,
zooxanthellate clionid sponges. If you look closely you will observe the
in/ex- current holes of the sponge and sponge tissue instead of live coral
polyps. If you can see the coral skeleton beneath the sponge, and are able to
identify it to genus or even species, include the affected area in your
estimate of “old death” and note “Cliona overgrowth” in the
corresponding Comments box.
"Recently dead" is defined as any non-living parts of the coral in which the corallite structures are either white and still intact or slightly eroded but identifiable to species. Recently dead skeletons may be covered by sediment or a thin layer of turf algae.
Note: In some cases circular or oblong
lesions or excavations caused by fish biting may result in destruction of the
corallites. If fish bites are identifiable and constitute part of the
mortality, include the affected area in your estimate of “recent death.”
Note: How to assess corals that
are detached from the substratum:
i. If it has recently fallen,
the length, height and % mortality should be measured as if it were still
upright; write “fallen” in comments box.
ii. A detached but wedged coral should be marked
as “wedged” in the comments section (as it is likely to remain in this position
for an extended period).
iii.
If it has been fallen for long enough to have reoriented to grow upward in its new position, the “new” maximum
length and maximum width should be measured, and the new outward-facing surface
used for calculating % mortality.
e. Scan over the surviving portions of the ENTIRE coral colony for any DISEASES and/or BLEACHED tissues present.
Characterize any DISEASES by the following color categories:
BB = Black band
WB = White band (Acropora only)
WS = White patches/white pox/patchy necrosis (Acropora only)
WP = White plague
YB = Yellow-band/yellow-blotch
RB = Red band
For more information about coral diseases see:
Bruckner (2002) Appendix II Coral Health and Mortality. Recognizing the signs of coral diseases and predators. Pp. 240-278 in P. Humann, ed., Reef Coral Identification. New World Publications, Inc.);
or one of the following web sites:
http://www.unep-wcmc.org/marine/coraldis/cd/index.htm
http://www.coral.noaa.gov/coral_disease/
Characterize any BLEACHED tissues as approximate severity of discoloration:
P = Pale (discoloration of coral tissue)
PB = Partly Bleached (patches of fully bleached or white tissue)
BL = Bleached (tissue is totally white, no zooxanthallae visible)
Many severely bleached corals are translucent, but you can still see the polyp tissues above the skeleton. Bleached tissues should not be included with the “recently dead” estimates.
Note:
It is important to be able to differentiate between tissues that are alive but
look white because they are bleached and white, recently dead skeletons.
f. If present, record in the appropriate column the presence of any damselfish algal gardens and the number of territorial gardening damselfish [Stegastes diencaeus (longfin), S. fuscus (dusky), S. planifrons (threespot), or S. variabilis (cocoa)] associated with the coral. (You may have to wait a couple of seconds to let the fish come back or pop-up from hiding after assessing the coral.) Ignore any herbivorous Microspathodon chrysurus (yellowtail), which are surveyed during the fish transects, and the planktivorous Stegastes partitus (bicolor).
g. Record any other sources of recent mortality that can still be unambiguously identified. Possibilities include sediments, storm damage, parrotfish bites, predation on the soft tissues by snails like Corallophilia abbreviata or the bristle worm Hermodice carunculata, various effects of adjacent benthic macroalgae or sediment-bound algal turfs, and any other spatial competitors (e.g., zoanthids like Palythoa caribeorum and Zoanthus spp., encrusting gorgonians as Briareum asbestinum and Erythropodium caribaeorum, tunicates like Trididemnum solidum and Didemnum vanderhorsti, or other stony corals).
How to assess large clusters or thickets in which colony
boundaries are not distinguishable:
Write “Point Counts” in the Comments box. Measure the total live coral
intercept length, maximum length, maximum width and maximum height for the
clump as a whole. Using the 1-m pole for scale, in the Comments box, record the
condition of the points at 10 cm intervals along the transect line as # live
(L), # recently dead (RD), and # old dead (OD) (i.e., 9L, 1RD, 3OD = nine
points that were alive, one that was recently dead and three that were old
dead). Record any signs of disease,
bleaching overgrowth, etc., for the clump as a whole.
5. As you advance along the transect measure, and write down, the cover under the line of each of the following:
Sand (only loose and deep enough to prevent coral larvae from settling);
Live Coral (for
colonies that are < 10 cm in length);
Crustose Coralline Algae;
Fleshy Macroalgae;
Calcareous Macroalgae;
Other Sessile Benthic Animals (i.e., any gorgonians, sponges, zoanthids, tunicates, etc.).
There is space on the datasheet for separate measurements as each is encountered; the totals for each category can be calculated after the dive. Do not measure the cover of algal turfs or “barren” areas of dead corals, rubble, hardbottoms or pavements.
Third pass:
6. Re-swim the line with the 25 X 25 cm quadrat and the 1-m pole. Starting at the 1-m mark, place the quadrat every two meters directly below the meter mark on the transect line.
a. For each quadrat, record each of the following:
Substratum–as pavement (pv), live coral (lc), dead coral (dc), rubble (rb) or sand (sn);
Fleshy Macroalgal Height–approximate their average canopy height with the plastic ruler;
Calcareous Macroalgal Height–approximate their average canopy height with the plastic ruler;
“Recruits”–the number of all small (up to 2 cm maximum diameter) stony corals (scleractinians and Millepora) in the quadrat. Identify any as you can to the genus or species level.
If there are no fleshy and/or calcareous macroalgae within the quadrat (as may occur when the 1-, 3-, 5-, 7- or 9- m mark is in the middle of a large, live coral or a sand patch), measure their average canopy heights at the closest point under the line in which they occur as you continue along the transect.
b. Maximum Reef Relief within a 1m radius of the 1-, 3-, 5-, 7-, and 9- m marks along the transect line, measure the height of the tallest coral or reef rock above the lowest point in the underlying substratum.
c. If you can identify the most common macroalgae in the quadrats, write their names in General Comments.
7. After you complete a transect, collect the line and haphazardly reset the next transect line, at least 2 m laterally away from its previous position. Remember to avoid other lines, and whenever possible, abrupt changes in slope, large areas of sand and rubble, and any other unusual reef features. Try to ensure that the transect lines are distributed around the SITE, and not concentrated close together.
8. Repeat the above steps for each transect.
9. Continue to reset transects in new positions until there are a minimum of 6 transects per SITE. Although appropriate sample sizes will depend on the variance in the local habitats, hence we cannot prescribe “a one size fits all protocol,” a minimum of 30 quadrats and 50 corals that are ≥10 cm should be assessed at each SITE
10. After surveying, enter your data into a copy of the provided AGRRA spreadsheet in Microsoft Excel. (Be sure to use a separate copy of the spreadsheet for every SITE.) Please check your data to verify its accuracy, then submit an electronic copy to the AGRRA database. Back up your own data regularly and store it in a safe place.
Note: Please examine carefully the example of the
datasheet below, and make sure you understand the instructions on how to fill
in all the cells in the form.
coral reef fishes
Method I: Belt transect
counts for defined species list.
1. For each transect, record the following information in the same fashion as explained in the coral section: your name (four letter code), date, time of start of transect, site name, site number, latitude and longitude, transect # on the UW datasheet.
2. Swim a 30-m transect line by first placing the weighted end of the line on the bottom (fix to some crevice or rock, so it won’t drag when the reel stops), and then swimming in a straight line while releasing it from the reel as you count the fish. This minimizes the disturbance to the fishes prior to their being counted. Periodically fixing on an object in the distance as you swim will help you swim in a straight line. (You can clip the transect tape to your weight belt to allow for easy release of the tape).
3. As you swim out the full 30 m transect line, count and record fish found within a visually estimated belt transect that is 2-m wide and extends up to the water surface. Use a 1-m wide T-bar to ensure accurate monitoring of the 2-m wide belt. Hold the T-bar ahead of you angled downward at about 45 degrees, and try to focus your gaze on the several meters of the transect ahead of the T-bar. Count only the species listed below and do not count juvenile parrotfishes or grunts that are less than 5 cm in total length. This list of species has been chosen to provide coverage of a number of the species most likely to be affected by human impacts, while preserving a relatively consistent search image to enhance the precision of transect data.
AGRRA Fishes–unless otherwise noted below, include every species within each of the following families:
Acanthuridae (Surgeonfish) ALL (e.g., Acanthurus bahianus, A. chirurgus, A. coeruleus);
Balistidae (Triggerfish) ONLY
Balistes vetula (queen triggerfish), B. capriscus (gray triggerfish), Melichthys
Chaetodontidae (Butterflyfish) ALL (e.g., Chaetodon capistratus);
Haemulidae (Grunt) ALL ≥ 5 cm (e.g., Haemulon flavolineatum, H. chrysargyreum, H. sciurus, H. plumieri);
Lutjanidae (Snapper) ALL (e.g., Lutjanus griseus, L. apodus, L. mahogoni, Ocyurus chrysurus);
Pomacanthidae (Angelfish) (e.g., Pomacanthus paru, P. arcuatus, Holocanthus tricolor);
Scaridae (Parrotfish) ALL ≥ 5 cm (e.g., Sparisoma viride, S. aurofrenatum, Scarus taeniopterus, S. vetula);
Serranidae (Groupers), ONLY Epinephelus spp. and Mycteroperca spp. (e.g., Epinephelus guttatus, E. fulvus, E. striatus, Mycteroperca bonaci).
Also count each of the following five species:
Bodianus rufus (Spanish hogfish)
Caranx rubber (Bar jack)
Lachnolaimus maximus (Hogfish)
Microspathodon chrysurus (Yellowtail damselfish)
Sphyraena barracuda (Barracuda)
4. Estimate the size of each fish with the 5-cm increments on the 1 m T-bar, and assign them to the following size categories: <5 cm (excepting acanthurids and scarids); 5-10 cm; 10-20 cm; 20-30 cm; 30-40 cm; >40 cm. Large groups of individuals of a species will be classified by attempting to put them into one or more size categories as necessary. By remembering to keep effort equivalent on all segments of the transect, you can limit the tendency to count all members of a school crossing the transect, instead of just those members which happen to be within the transect as counting of that segment takes place.
Note: Sample the transect belt giving
uniform attention to each successive 2-m segment. This requires swimming at a
more or less constant rate, while looking consistently about 2 m ahead of your
current position. You may pause while recording data, and then start swimming
again. It is important to swim in a uniform manner. A speed that covers each
30-m transect in 6-8 minutes should be attempted. High densities of counted
species will slow this rate in some cases. Fish observers should be trained to
estimate fish lengths by using consistency training methods both on land and
underwater.
5. When you reach the end of the transect line, stop the survey and recoil the transect tape.
6. Continue conducting haphazardly-positioned 30 m transects at least 5 m laterally away from the previous position. Repeat the above steps for each transect.
7. Conduct a
minimum of ten (10) transects at each site.
Modifications: Some workers may want to census other species of fish. This is encouraged, provided that these other species are counted on a separate pass in the same area, after the AGRRA run. Otherwise the census method is substantially changed, and your data may not be directly cross-comparable with other AGRRA assessments.
Method II. Rover Diver census
After finishing the belt transects (or concurrently depending on the number of surveyors), conduct a roving diver census of all species of fishes following the methodology of Reef Environmental Education Foundation (REEF) (http://www.reef.org/) and briefly explained below.
1. The Rover diver census is conducted in the same general area as the belt transects are set.
2. Swim around the reef site for approximately 30 minutes (preferably 45-60 min) and record ALL fish species observed. Use all knowledge you have of fish habits, and search under overhangs, in caves, and so on. The objective is to find the maximum number of species that you can in during your search time.
3. Estimate the density of each species by using logarithmic categories: Single (1fish), Few (2-10 fishes), Many (11-100 fishes), or Abundant (>100 fishes).
4. Record your observations on the standardized REEF data entry sheet.
5. Submit data to REEF database.
OPTIONAL COMPONENTS
Several other
useful assessments may be easily integrated into the core portion of the protocol
given above. These optional components, while not part of the core methods, can
yield additional information that may lead to a better understanding of the
condition of a reef. These optional components include coral recruitment
(additional sampling effort) and fish bites.
Coral Recruitment: Coral recruitment is an important indicator of a reef’s regeneration potential and is now incorporated into the standard AGGRA protocol. If you would like to increase your sample size of coral recruits, we suggest the following:
Method:
1. After you have completed your 6 standard benthic transects (you may plan an extra dive at the same location for this procedure), lay a set of 10 haphazard transects, placing the 25 x 25 cm quadrat on the substratum right below each of the odd meter marks (1,3,5,7,9). The objective is to assess 50 extra quadrats for a minimum sample size of 80 quadrats (30 from the first 6 transects + 50 from the extra 10 transects), which equals an area of 5 sq. meters.
2. Count all small (maximum diameter 2 cm), stony corals (scleractinians and Millepora) that you can see within the 25 X 25 cm quadrats.
3. Whenever possible, record their scientific names at least to the level of genus.
4. Try to repeat for a total of at least 80 quadrats (an overall sample of 5 square meters of reef surface).
Note: Proper training and good eyesight (or
corrective lenses) are essential to accurately detect the presence of small
corals due to their inconspicuous size and nature.
Herbivory
The objective of the Fish Bite Method (Steneck 1985) is to gauge the effect of herbivorous fishes on algal composition by quantifying their level of herbivory. Fish herbivory is assessed by counting the observed number of bites per square meter of different guilds of herbivorous fishes, which are categorized as:
Scrapers = Scaridae (parrotfish)
Browsers = Acanthuridae (surgeonfish), Microspathodon chrysurus (yellowtail damselfish)
Non-denuders = other Pomacentridae (damselfish) but not Stegastes partitus (bicolor damselfish).
Method:
1. Use the 1 m stick in conjunction with natural landmarks on the reef surface (e.g., a small coral or gorgonian) to haphazardly delineate an area that is approximately 1 m square and representative of the benthic cover on the reef substratum. (Please do not place a meter quadrat to mark your observation area, as some fish are particularly prone to biting novel objects placed within their feeding territories).
2. Back off as far as you can while still seeing the meter square area. Watch for 5 minutes. Record the depth, time of day, and number of bites from all species of fishes in the three guilds listed above (identify them to species as best as you can). Repeat for a total of 5 squares (and ~25 minutes of observation).
Note:
You must be able to distinguish (a) juvenile scarids from other fishes
with similar stripes, such as acanthurids and labrids (wrasses- which only look
as though they are biting algae as they search for amphipods to eat) and (b) yellowtail
damselfish (which are browsers) from other species of damselfish (that
cultivate algal gardens). Be sure to remember to record the time of day since
fish activity varies temporally.
Appendix
How to paint the 1 m sticks:
Cut ½” PVC water pipe into a
few 1m long segments.
To build a painting template
(Figure 1):
Cut a ¾” PVC water pipe into
a 1 m segment, and then cut it in half longitudinally.
Cut ¾” PVC pipe into five 10
cm segments and glue them to the 1 m half pipe segment, spaced exactly 10 cm
from each other.
Slide in one of the 1 m ½”
segments through the template, matching both ends, one must be covered and the
other uncovered.
Apply spray paint to the ½”
pipe rotating inside of the template to cover the whole circumference of the
pipe.
Let it dry a bit and start
again from #3.
Figure 1. Template for painting 1 m sticks
AGRRA kit for Benthic Survey
Views of the transect line
wrapped tightly around the quadrat, holding the 1 m stick and the clipboard
(with rubber bands for securing the benthic datasheets to the clipboard).
(a) front view
(b) back view
(c)
Details of the quadrat: note the holes on the corners to let the air and water out,
example of the rubber band leash for rulers and pencils (If available, we
recommend PaperMate ® Sharpwriter
mechanical pencils).
AGRRA
kit for fish surveys.
(a) T-bar with a clip and a
slit to hold the half/letter size board, and rubber bands to secure the UW
datasheets.
(b) Details on the handle and board-fastening features.
Fish Memory Cues (common AGRRA
species)
Angelfishes
21 Queen Angelfish
The Queen has a crown (dark blue spot on forehead ringed with bright
blue).
23 French Angelfish
This fashionable French beauty is dressed in classic black (with gold
highlights).
25 Gray Angelfish
As its name implies, the Gray is gray to grayish brown.
25 Rock Beauty
This little beauty is yellow and black. The juvenile is bright yellow
with a small black spot (ringed in blue). The black spreads as the fish grows
covering most of the fish as an adult.
Butterflyfishes
29 Banded Butterflyfish
White with black bands (thick diagonal black markings).
29 Foureye Butterflyfish
Large false “eyespot” on tail.
31 Spotfin Butterflyfish
Small black spot on the rear of the bright yellow dorsal fin.
31 Reef Butterflyfish
Uncommon – identification by process of elimination (no good memory
clue).
33 Longsnout Butterflyfish
Tiny fish with long pointy snout (as name implies). Usually found deep.
Surgeonfishes
33 Blue Tang
Blue with contrasting yellow “tang” (spine on base of tail). Juveniles
change from all yellow to combination of yellow and blue to all blue as adult.
35 Ocean Surgeonfish
Clear pectoral fin – think “
37 Doctorfish
Dark pigmented leading edge of pectoral fin – think “Dark Doctor”.
Silvery (Miscellaneous)
45 Bar Jack
Most common jack with black and blue “crowBAR” along back and onto lower
tail fin.
65 Great Barracuda
Large, silvery, toothy torpedo. Most divers (and non-divers) know this
species.
Grunts
93 French Grunt
Diagonal gold markings like the gold braids worn on a French General’s
uniform.
93 Bluestriped Grunt
Blue horizontal stripes over yellow body. If pale in shallow water, black
rear dorsal and tail fin are good ID cue.
93 Smallmouth Grunt
Small grunt – (Small Mouth). Silvery fish with horizontal yellow lines and
yellow fins.
95 White Grunt
All fins white. Body checkered pattern of pearly white, blue & yellow
formed by scales. Thin stripes only on head.
95 Caesar Grunt
Silvery with thin yellow lines like raw egg drizzled over a Caesar salad.
Dusky rear dorsal, anal, and tail fins like the dusty feet of Caesar’s army.
97 Tomtate
Whitish fish with two thin yellow lines (one midbody through eye, the
other on back). Usually a black spot at base of tail. Think of a TomTom (a
small drum) with the two yellow lines as drum sticks.
97 Cottonwick
Black line from the snout through the eye fades as it reaches the tail.
Think of the black cotton wick of a candle. Usually have a black diagonal
stripe that runs along the back and onto the tail.
99 Spanish Grunt
Large grunt with horizontal black lines and a yellow saddle on the base
of the tail. Think of the fried egg in a Spanish omelette.
101 Sailors Choice
Silvery gray fish with distinctive black spots on scales covering the
body; gold ring encircles the eye. Think pirates (who were sailors) with the
black spots as rows of waves and the gold ring as an golden earring.
107 Porkfish
Two black diagonal bands on head (one through eye and the other just
behind the gills). For pork, think of the bands as two strips of overcooked
bacon.
109 Black
The large black patch on the side of this fish makes the Black Margate
easy to remember.
109 White
About the size of a Black Margate but without the black patch. Very steep
forehead with high back profile. Eye is tiny with white iris.
Snappers
111 Mutton Snapper
This species is easiest to ID if you know that its scientific name is analis since it is the only snapper with
a pointed (not rounded) anal fin. It usually has a small black spot on the back
(“the button on the Mutton”) which we can use to remember its common name.
111 Cubera Snapper
This is the largest of the snappers (up to 3’), usually solitary, and
often with pale bars across back.
113 Gray Snapper
Gray with no distinguishing features other than a dark diagonal band that
occasionally runs from lip across eye.
115 Dog Snapper
Has “teardrop stains” below eye. For the girls we say the fish is crying
because it lost its dog; for the guys we go for the more macho memory cue of
“dog tags”.
115 Mahogany Snapper
Silvery white fish with “Mahogany” red margin on tail; sometimes reddish
tinge on body or other fins.
117 Lane Snapper
Though sometimes faint, this fish has yellow “lane” markers (think
highway) along it’s body. It may have a small black spot just below the rear
dorsal.
119 Yellowtail Snapper
Bright yellow midbody stripe continues onto yellow tail. Feed in the
water column high above reefs.
121 Schoolmaster
Large silvery white fish with all yellow fins.
Damselfishes
133 Yellowtail Damselfish
The only damselfish we need to know and one of the easiest to remember as
it has a yellow tail. Juveniles are bright blue with brilliant blue spots. The
tail is translucent on very young juveniles.
Groupers/Sea Basses
153
The black saddle is the easiest way to ID this fish. Think “Ride the
Nassau Grouper back to the
157 Graysby
Most common of the smaller groupers. Grayish brown with 3-5 pale or dark
spots along back along base of dorsal fin. If you want you can think of
“buttons down the vest of the Great Gatsby”.
159 Red Hind
Reddish spots over a lighter background rear fins (rear dorsal, tail, and
anal) edged in black. Think “RED with a black beHIND”.
159 Rock Hind
Have a black saddle (and usually additional black blotches along back
under the dorsal fin). Think of these spots as “rocks”.
161 Coney
This variable species can be reddish brown, bicolor (upper dark lower
pale), or a brilliant yellow so color is not a good ID cue. The body is usually
covered with tiny blue spots. One constant is that it has two spots on the
lower lip and two on the base of the tail.
163 Black Grouper
Blotches on back squarish. Think “Black Bricks” or “Black Blocks”.
165 Tiger Grouper
Have “tiger-strips” across back. Also have some pretty impressive canine
teeth.
167 Yellowmouth Grouper
Corners of the mouth a distinctive yellow. Margins of pectoral fins pale.
169 Yellowfin Grouper
Margins of pectoral fins yellow. Blotches on back are more oval and not squarish
like the Black Grouper.
Parrotfishes
195 Blue Parrotfish
Adults are blue with no markings. Juveniles have a yellow wash on the
head.
195
Dark navy (“
197 Rainbow Parrotfish
“Rainbow” colored with orangish head and tail and bright green rear body.
197 Queen Parrotfish
TP: Queen has a moustache and beard (blue/green markings around mouth).
IP: Black and white like a chess board.
199 Stoplight Parrotfish
JP & IP: Bright red belly (like a stoplight).
TP: Small yellow spot at top of gill cover. (Like the yellow light in a
middle of a traffic light?)
199 Princess Parrotfish
TP: Tail bordered with pink. Think “Pink Princess”.
JP: Looks like the Striped Parrotfish juvenile but doesn’t have a gold
nose. Think “The Princess has no gold”.
201 Striped Parrotfish
TP: The tail is not bordered in blue (not pink like the “Pink Princess”).
JP: The Princess Parrotfish may be royalty but it is the Striped that has
the gold (on its nose).
203 Redband Parrotfish
Exceedingly variable parrotfish. Only the TP have the namesake “redband”
across the cheek. In all of the other JP/IP color variations, look for the
white spot (saddle) on the base of the tail.
205 Redtail Parrotfish
TP: Red crescent in the middle of the tail.
IP: Red tail (and body) – mostly reddish gray can be pale.
205 Redfin Parrotfish
Also known as Yellowtail Parrotfish. Name comes from small reddish spot
at base of pectoral fin but yellow tail is usually more visible. I always
remember this fish by both of its names when I see it and think “yellowtail
=> redfin”.
209 Greenblotch Parrotfish
Tiny parrotfish named Greenblotch for the green blotch on the side of the
TP. The JP/IP are usually red to yellowish red. All phases have a bright
yellow-gold to red iris.
Hogfishes
213 Hogfish
The spiky front dorsal finare like the bristles on the back of a
razorback hog.
213 Spanish Hogfish
Think of the purple area across the top of the body as stain from a
bottle of Spanish wine.
Leatherjackets (Triggerfishes
& Filefishes)
395 Queen Triggerfish
The Queen is long eyelashes (the black lines radiating from the eyes) but
like the Queen Parrotfish, she has a blue moustache.
397 Ocean Triggerfish
Uniformly gray and usually swimming high in the water column. Has a black
spot at base of pectoral fin.
399 Black Durgon
Usually black overall (with pale bluish white lines along base of dorsal
and anal fins. Can have a bluish or greenish cast.
401 Whitespotted Filefish
Large orange, brown and gray colored fish often with large white spots.
Commonly seen in pairs with one fish showing spots, the other without. Pair of
orange spines at tail base.
403 Orangespotted Filefish
Dusky brownish color with small orange spots (more intense on back and
toward tail). Small white saddle on tail is distinctive.
New Datasheets (fish & coral)
| |
|
Robert N. Ginsburg Atlantic and Gulf Rapid Reef Assessment MGG-RSMAS, University of Miami 4600 Rickenbacker Causeway Miami, FL 33149 USA |
Telephone: (305) 421-4664 Email: info@agrra.org Send data to: data@agrra.org URL: http://www.agrra.org |
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