Methods for Monitoring Corals and Crustose Coralline 

Algae to Quantify In-Situ Calcification Rates

U.S. Department of the Interior

U.S. Geological Survey

Open-File Report 2013–1159
Cover photograph: Calcification-monitoring station, Molasses Reef, Florida 

Keys National Marine Sanctuary. Photograph by Ilsa Kuffner, November 2012. 
Methods for Monitoring Corals and 

Crustose Coralline Algae to Quantify 

In-Situ Calcification Rates

By Jennifer M. Morrison, Ilsa B. Kuffner, and T. Don Hickey

Open-File Report 2013–1159

U.S. Department of the Interior

U.S. Geological Survey
U.S. Department of the Interior

SALLY JEWELL, Secretary

U.S. Geological Survey

Suzette M. Kimball, Acting Director

U.S. Geological Survey, Reston, Virginia: 2013

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Suggested citation:

Morrison, J.M., Kuffner, I.B., and Hickey, T.D., 2013, Methods for monitoring corals and crustose coralline algae to 

quantify in-situ calcification rates: U.S. Geological Survey Open-File Report 2013–1159, 11 p., http://pubs.usgs.gov/

of/2013/1159/.
iii

Acknowledgments

The development of this calcification monitoring network was principally funded by the 

U.S. Geological Survey (USGS) Coastal & Marine Geology Program, with supplementary funds 

from the USGS Terrestrial, Freshwater, and Marine Ecosystems Program and grants from the 

USGS Global Change Research Fund and the Department of Interior Southeast Climate Science 

Center. For help in setting up and maintaining the calcification stations, we thank A. Brame, J. 

Sanford, C. Reich, C. Reynolds, and C. Williams. We thank K. Ludwig for engineering the tilesaw modifications and K. Hargrave for the illustration in figure 1. We thank C. Reich and 

C. Williams for their constructive comments on the manuscript.

We express our gratitude to the National Park Service and the Florida Keys National Marine 

Sanctuary for permitting the monitoring network under scientific permits DRTO-2009-SCI-0009, 

DRTO-2011-SCI-0004, BISC-2009-SCI-0019, BISC-2010-SCI-0035, BISC-2011-SCI-0025FKNMS2008-062A, and FKNMS-2010-122.
iv

Contents

Abstract...........................................................................................................................................................1

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

Methods...........................................................................................................................................................2

Station Setup and Design....................................................................................................................2

Corals......................................................................................................................................................3

Crustose Coralline Algae.....................................................................................................................5

Monitoring and Sampling ....................................................................................................................6

Results and Discussion.................................................................................................................................7

Summary..........................................................................................................................................................9

References Cited............................................................................................................................................9

Figures

1. Exploded diagram of a coral and crustose coralline algal calcification station, 

including all fasteners and attachments ..................................................................................3

2. Photographs showing mounting, staining, and weighing of coral colonies.......................4

3. Photographs showing monitoring and sampling methods ....................................................6

4. Photographs showing plastic “pear” neck tags, used in the livestock industry, 

used here as algal recruitment tiles..........................................................................................7

Table

1. Materials used to construct each calcification station. All fasteners were marinegrade stainless steel (alloy 316).................................................................................................5
v

Conversion Factors

Inch/Pound to SI

Multiply By To obtain

Length

inch (in.) 2.54 centimeter (cm)

inch (in.) 25.4 millimeter (mm)

foot (ft) 0.3048 meter (m)

mile (mi) 1.609 kilometer (km)

Volume

gallon (gal) 3.785 liter (L) 

Temperature in degrees Celsius (°C) may be converted to degrees Fahrenheit (°F) as follows:

°F=(1.8×°C)+32

Temperature in degrees Fahrenheit (°F) may be converted to degrees Celsius (°C) as follows:

°C=(°F-32)/1.8

Abstract

The potential effect of global climate change on calcifying marine organisms, such as scleractinian (reef-building) 

corals, is becoming increasingly evident. Understanding 

the process of coral calcification and establishing baseline 

calcification rates are necessary to detect future changes 

in growth resulting from climate change or other stressors. 

Here we describe the methods used to establish a network of 

calcification-monitoring stations along the outer Florida Keys 

Reef Tract in 2009. In addition to detailing the initial setup 

and periodic monitoring of calcification stations, we discuss 

the utility and success of our design and offer suggestions for 

future deployments. Stations were designed such that whole 

coral colonies were securely attached to fixed apparati (n = 10 

at each site) on the seafloor but also could be easily removed 

and reattached as needed for periodic weighing. Corals were 

weighed every 6 months, using the buoyant weight technique, 

to determine calcification rates in situ. Sites were visited 

in May and November to obtain winter and summer rates, 

respectively, and identify seasonal patterns in calcification. 

Calcification rates of the crustose coralline algal community 

also were measured by affixing commercially available plastic 

tiles, deployed vertically, at each station. Colonization by 

invertebrates and fleshy algae on the tiles was low, indicating 

relative specificity for the crustose coralline algal community. 

We also describe a new, nonlethal technique for sampling 

the corals, used following the completion of the monitoring 

period, in which two slabs were obtained from the center of 

each colony. Sampled corals were reattached to the seafloor, 

and most corals had completely recovered within 6 months. 

The station design and sampling methods described herein 

provide an effective approach to assessing coral and crustose 

coralline algal calcification rates across time and space, offering the ability to quantify the potential effects of ocean warming and acidification on calcification processes.

Introduction

The calcium carbonate skeletons produced by scleractinian corals constitute the foundation of coral reef ecosystems, 

providing habitat and protection for a diverse array of marine 

organisms. Coral reefs also offer immeasurable value to 

humans through tourism, fisheries, and the protection of coasts 

from erosion. In the last few decades, corals have faced an 

increasing number of local, chronic stressors (such as disease, 

coastal development and sedimentation, eutrophication, and 

overfishing) as well as extreme temperature anomaly events, 

the effects of which are alarmingly apparent in declining coral 

populations worldwide (De’ath and others, 2012; Costa Jr. and 

others, 2008; Edinger and others, 1998; Fabricius, 2005; Gardner and others, 2003; Jackson and others, 2001; Reopanichkul 

and others, 2009; Hughes, 1994; Wolanski and others, 2009; 

Aronson and others, 2002; Berkelmans and Oliver, 1999; 

Eakin and others, 2010). Moving into the future, increased 

frequency and intensity of warm-water-induced bleaching 

events, associated with ocean warming, and decreased carbonate ion concentrations (ocean acidification) represent the two 

most critical long-term threats to coral reefs (Hoegh-Guldberg, 

1999; Hoegh-Guldberg and others, 2007).

Reef-building corals are especially susceptible to ocean 

warming and acidification because of the narrow water temperature range and high carbonate ion concentration needed 

for optimal skeletal production (Cantin and others, 2010; Carricart-Ganivet, 2004; Kleypas and Langdon, 2006; Hughes and 

others, 2003; Veron and others, 2009; Andersson and Gledhill, 

2013; De’ath and others, 2009). It is necessary to document 

current patterns in calcification to detect and quantify future 

changes, resulting from natural or anthropogenic stressors. 

Estimates of calcification rates of individual coral colonies are 

most commonly obtained through measurements of skeletal 

extension rates. To determine linear extension, cores are 

collected from live corals, sliced parallel to the main growth 

axis, and x-rayed (Knutson and others, 1972; Hudson, 1981). 

The distance between adjacent high-density bands in the 

skeleton (annual linear extension), visible in the x-ray image, 

is measured and used to extrapolate the calcification rate by 

multiplying the linear extension rate by the average bulk skeletal density. There are several inherent limitations and issues 

with using this method, as demonstrated by the large amount 

of variability in the extension rates reported in such studies 

(Carricart-Ganivet, 2011). Measuring calcification directly, as 

opposed to calculating it using linear extension, may provide 

less variable data. In addition, direct measurements enable us 

Methods for Monitoring Corals and Crustose Coralline 

Algae to Quantify In-Situ Calcification Rates

By Jennifer M. Morrison, Ilsa B. Kuffner, and T. Don Hickey
2 Methods for Monitoring Corals and Crustose Coralline Algae to Quantify In-Situ Calcification Rates

to assess sub-annual patterns in calcification rates and make 

more direct inferences regarding the environmental variables 

driving calcification processes.

Whereas an abundance of studies have focused on establishing growth rates of reef-building corals, there are few studies reporting calcification rates of the crustose coralline algal 

(CCA) community. Though CCA may not calcify as quickly 

as reef-building corals (Adey and Vassar, 1975; Kuffner and 

others, 2013), their benthic cover far exceeds that of corals, in 

terms of geographical expanse and the amount of substratum 

covered in the photic zone (Steneck, 1986). Crustose coralline algae are ubiquitous on reefs and play a critical role in the 

development and stabilization/cementation of reef structures 

(Tierney and Johnson, 2012; Björk and others, 1995), the production of carbonate sediments (Adey and Macintyre, 1973), 

and the attraction of coral larvae and other invertebrates for 

settlement (Morse and others, 1988; Whalan and others, 2012). 

The importance of CCA, both ecologically and structurally, to 

coral reef ecosystems, is reason enough to merit further study. 

Furthermore, previous studies have demonstrated the vulnerability of CCA to simulated ocean acidification conditions 

(Kuffner and others, 2008; Hall-Spencer and others, 2008; 

Anthony and others, 2008), suggesting that monitoring CCA 

will become increasingly important in the future and could 

serve as an indicator of the effect and regional severity of 

ocean acidification.

This report discusses in detail the methods and materials 

used to construct and maintain a network of calcificationmonitoring stations along the outer Florida Keys Reef Tract 

(FKRT), which has proven successful in documenting spatial 

and temporal differences in coral and CCA calcification rates 

(Kuffner and others, 2013). This monitoring network is the 

first of its kind to systematically measure in-situ calcification 

rates of whole coral colonies and CCA in a reef-crest environment. Corals were monitored over a 2-year period, during 

which time they were removed for weighing every 6 months. 

Algal recruitment tiles were deployed every 6 months to 

measure net CCA accretion. We also introduce in this report a 

nonlethal method of sampling coral colonies, which provides 

a permanent, whole-colony growth record of the corals for 

additional analyses (such as measurements of linear extension, 

bio-erosion, stable isotopes, and trace elements) and preserves 

the remaining live coral for placement back on the reef.

Methods

Station Setup and Design

Stations were designed specifically for the purpose of 

regular, long-term (multi-year) monitoring of coral and CCA 

calcification rates. We chose to use small, intact colonies of 

coral rather than nubbins or fragments (Cook and others, 

2002), to capture growth true to whole-colony morphology. We also used independent blocks instead of racks for 

mounting the corals, for the sake of statistical power (each 

block represented a statistically independent replicate). 

This study was further set apart from other in-situ calcification 

studies by the fact that, rather than studying coral growth at 

more accessible, inshore sites to infer growth at offshore 

reefs, monitoring took place at sites along the outer reef 

tract. Specific criteria for site selection included habitat 

type (spur-and-groove or low-relief hard bottom), depth (12 

to 18 feet), and the availability of oceanographic and meteorological data. For the first round of monitoring, we chose 

to study the massive, reef-building coral Siderastrea siderea

because of its relative abundance along the FKRT and its resilience (Muthiga and Szmant, 1987; Lirman and Fong, 2007; 

Colella and others, 2012). The station design and monitoring 

methodology, however, could easily accommodate small colonies of any coral species.

A single station consisted of an 8 x 8 x 8-inch concrete 

block (“half block”) with a small, whole coral colony and 

an algal recruitment tile attached (fig. 1). We used concrete 

blocks as the basis of the calcification stations because they 

are commercially available everywhere and are not susceptible 

to corrosion. Marine-grade stainless steel (alloy 316) hardware 

was used because of its corrosion resistance, and all fasteners 

were of the same alloy to prevent electrolytic corrosion. All 

fasteners had a 1/4-inch screw diameter and 20 threads per 

inch (“1/4 - 20”). Prior to installation, a single, approximately 1

/4-inch hole (slightly larger than the cap screw diameter for 

easy removal of the coral for weighing) was drilled into the 

top-facing side of each concrete block for mounting the coral 

colony; two 1/4-inch holes were drilled diagonally across the 

bottom-facing side of the block, roughly 4 inches apart, to 

allow the block to be secured to the substratum; and two 

1

/4-inch holes were drilled on one side of the block, approximately 1 inch apart, for mounting the algal recruitment apparatus (see fig. 1 for approximate hole locations). Stations were 

installed in areas of low-relief, hard-bottom habitat, where 

other corals were presently growing and where there was relatively flat, secure substratum available for block installation.

To install each station, two holes, spaced roughly 4 inches 

apart [using a template made of polyvinyl chloride (PVC) 

sheeting], were drilled 2 to 3 inches deep into the substratum 

using a pneumatic drill (with a 1/2-inch masonry bit), powered 

by an air compressor located onboard a two-point-anchored 

vessel. The substratum surrounding the holes was then scraped 

with a rock chisel (to flatten the surface and remove turf algae) 

and brushed with a plastic, stiff-bristled brush (to remove sediment and other loose material), to ensure maximum adhesion 

of the epoxy to the seafloor. The holes were filled with All-Fix 

underwater epoxy (Cir-Cut Corporation, Philadelphia, Pennsylvania, U.S.A.), and a 6-inch-long threaded rod was inserted 

in each epoxy-filled hole, with care not to get any epoxy in 

the threads on the upper half of the rod. Additional epoxy was 

mounded around the base of the rods and on the substratum in 

between, to provide further support and adhesion for the rods 

and blocks, respectively. As soon as each pair of rods was in 

place (while the epoxy was still wet), the concrete block was 
Figure 1. Exploded diagram of a coral and crustose coralline algal calcification station, including all fasteners and 

attachments. Labels correspond with those listed in table 1.

Methods 3

lowered onto the rods (by lining up the holes on the bottom 

of the block) carefully, so as not to disturb the newly epoxied 

rods. Once the block was set, epoxy was added underneath 

the sides of the block as needed, to further stabilize the block 

on uneven substratum. Ten stations were installed at each site, 

spaced roughly 6 to 12 feet from each other.

Corals

While the epoxy cured, coral colonies were collected 

from a nearby site of similar depth and habitat type (Kuffner 

and others, 2013). Colonies were chosen that were hemispherically shaped (as opposed to encrusting), within the target 

size range (3- to 4-inch diameter), and lacking any visible 

signs of disease or clionid sponge infestation. Selected corals 

were removed from the reef by hammering (with a mallet) a 

rock chisel around the bottom of the colony. Once free from 

the substratum, a hammer and chisel were used to flatten the 

underside of the coral (if needed) and a wire brush was used 

to remove any loose debris, algae, or invertebrates. Epoxy 

was spread onto the bottom (chiseled side) of the coral, using 

excess putty to fill in any holes or crevices. The coral was then 

positioned on a round, gray PVC disc (3- or 4-inch diameter, 

depending on the size of the coral) that was roughened on 

one side with sandpaper and had a 1/4-inch hole drilled in the 

center and a 3-inch-long hex head cap screw placed in the 

hole (figs. 1 and 2A). The coral and roughened side of the disc 

were pushed together firmly to create a strong seal (fig. 2A). 

A unique, three-digit number was pre-etched onto the bottom 

side of each disc and was used to identify the corals throughout the study.

Once the coral colonies were collected and mounted onto 

PVC discs (n=10 for each site), they were transported back to 

the dock to be weighed and stained. Ten 5-gallon buckets were 

modified, prior to the first installation trip, for transporting and 

staining the corals. Each bucket was fitted with a plastic flower 
Figure 2. Mounting, staining, and weighing of coral colonies. A, Coral colony epoxied to the plastic disc and cap 

screw. B, Coral colony in a transport bucket. C, Coral colonies being stained with alizarin red S. D, Coral colony being 

weighed using an under-loading balance.

4 Methods for Monitoring Corals and Crustose Coralline Algae to Quantify In-Situ Calcification Rates

pot that had a 1/4-inch hole drilled in the bottom, three 1/4-inch 

holes drilled into the outer rim, and a roughly 4-inch hole cut 

out of the side. The outer rim of the flower pot was attached to 

the bottom of each bucket using nylon 1/4 - 20 bolts, inserted 

through 1/4-inch holes that were drilled into the bucket and 

the holes in the rim of the pot and secured with nylon 1/4 - 20 

hex nuts (fig. 2B). The flower pot and bolt connections were 

reinforced with hot glue. The buckets were then filled with 

seawater, and collected coral colonies were placed in the buckets, with the attached cap screw positioned through the 1

/4-inch 

hole in the bottom of the flower pot, securing the coral in an 

upright position (fig. 2B). To prevent heat stress in the corals 

while they were on the boat, the temperature of the water in 

the buckets was closely monitored so that it did not exceed 

30 °C. If the temperature neared this threshold, a shade cover 

was laid over the buckets and, if necessary, cool seawater was 

added.

At the dock, pre-weighed aliquots of alizarin red S 

(15 mg/L) were added to the seawater in each transport 

bucket. The corals were kept in the dye for 4 hours during the 

period surrounding the solar zenith (approximately 11 AM to 

3 PM Eastern Daylight Time), maximizing the incorporation 

of the stain into the coral skeleton. Staining the surface of the 

coral skeleton provided a time stamp for use in later growth 

measurements. For the duration of the staining period, the 

buckets were suspended in the ocean by buoyant rings (swim 

A B

C D
Table 1. Materials used to construct each calcification station. All fasteners were marine-grade stainless steel (alloy 316). 

[Dimension abbreviations: diam, diameter; in, inches; cm, centimeters; tpi, threads per inch; lg, length; mm, millimeters; p, pitch (distance between threads); t, 

thickness; w, width; ht, height; id, inner diameter; od, outer diameter]

Label Description Quantity Standard size Approximate metric equivalent

A Whole coral colony 1 3 – 4 in (diam) 7.5 – 10 cm (diam)

B Underwater epoxy

C Hex head cap screw, fully threaded 3 1⁄4 in (diam) – 20 (tpi) x 3 in (lg) 6 mm (diam) x 1 mm (p) x 70 mm (lg)

D Gray polyvinyl chloride (PVC) disc 1 3 or 4 in (diam) x 7⁄16 in (t) 7.5 or 10 cm (diam) x 11 mm (t)

E Concrete block (“half block”) 1 8 (lg) x 8 (w) x 8 in (ht) 20 (lg) x 20 (w) x 20 cm (ht)

F Fender washer 4 1⁄4 in (id) x 11⁄4 in (od) 6 mm (id) x 18 mm (od)

G Wing nut 1 1⁄4 in (id) – 20 (tpi) 6 mm (id) x 1 mm (p)

H Machine screw hex nut 8 1⁄4 in (id) – 20 (tpi) 6 mm (id) x 1 mm (p)

I Flat washer 2 1⁄4 in (id) x 9⁄16 in (od) 6 mm (id) x 18 mm (od)

J Blue “pear” livestock tag with station number 1 31⁄2 in (ht) 9 cm (ht)

K White nylon rod 1 7⁄8 in (diam) x 4 in (lg) 22 mm (diam) x 10 cm (lg)

L Zip tie (“miniature”) 2 4 in (lg) 10 cm (lg)

M Rubber o-ring (“AS568-211”) 2 1⁄8 in (t) x 13⁄16 in (id) 3.53 mm (t) x 20.22 mm (id)

N White “pear” livestock neck tag 1 31⁄2 in (ht) 9 cm (ht)

O Threaded rod 2 1⁄4 in (diam) – 20 (tpi) x 6 in (lg) 6 mm (diam) x 1 mm (p) x 15 cm (lg)

Methods 5

noodles), secured around the rim of each bucket, to maintain 

ambient seawater temperature inside the buckets (fig. 2C).

During the staining period, corals also were weighed 

using the buoyant weight technique (Jokiel and others, 1978). 

The weighing apparatus consisted of a heavy duty plastic 

bin, filled to a mark (at about 2/

3

 full) with seawater, and a 

plastic cooler, attached to the top of one half of the lid 

(fig. 2D). A battery-operated balance with an under-loading 

hook was placed inside the cooler, and a braided MicroDyneema FireLine® fishing line (Berkley®, Spirit Lake, Iowa, 

U.S.A.) was hung with a slip knot from the balance hook and 

the line passed through aligning 1-inch holes that had been 

cut through the bottom of the cooler and the plastic bin lid. 

A stainless steel washer was tied to the other end of the line 

so the corals could be attached for weighing. With the balance zeroed, the cap screw on the bottom of the coral was slid 

through the hole in the washer. The coral was gently lowered 

into the seawater in the bin until the line was taut (the weight 

of the coral provided enough tension between the threaded 

rod and the vertical washer to hold the coral in place without a 

fastener), making sure the coral was not resting on the bottom 

of the bin. The other one-half of the lid was then placed on the 

bin to minimize wind movement over the water, the balance 

was allowed time to settle, and the weight was recorded for 

each coral. The “bird’s eye” length and width dimensions were 

measured with calipers and used to calculate the planar surface 

area of the coral (using the formula for the area of an ellipse). 

The presence of any boring organisms on the surface of the 

colony also was noted before returning it to the transport 

bucket. The change in buoyant mass between weighing intervals was converted to dry mass, using a seawater density of 

1.02 g/cm3 and an aragonite density of 2.93 g/cm3 (see Jokiel 

and others, 1978 for conversion equations), and normalized to 

the 2-dimensional surface area of each coral.

When the epoxy had finished curing (at least 24 hours 

later) and the rods were firmly embedded in the reef, the 

concrete blocks were fastened down using a fender washer 

and two machine screw hex nuts on each rod (fig. 1, table 1). 

An identifying tag, with the station number (different from 

the coral number), was fixed between two washers on one of 

the rods (fig. 1, table 1). With the block secured to the substratum, the coral was then mounted to the block by inserting 

the attached cap screw through the hole in the top of the block 

and fastening it with a fender washer and wing nut (figs. 1 and 

3A, table 1). This allowed for secure attachment of the coral to 

the block as well as the ability to remove the coral easily for 

periodic weighing and staining.

Crustose Coralline Algae

To measure accretion rates of the CCA community, algal 

recruitment tiles were attached, in a vertical position, to the 

side of each block (fig. 3A). The recruitment tile, a white nylon 

“pear” livestock neck tag (National Band and Tag Co., Newport, Kentucky, U.S.A.), was fitted onto a 4-inch-long nylon 

rod with two pre-drilled 1⁄4-inch holes in the side (fig. 1, 

table 1). Commercially available tiles (fig. 4A) were used for 

this study because they were relatively inexpensive, did not 

require any preparation or cutting, and did not necessitate dissolving the samples to obtain dry mass, thereby allowing for 

sample archival. Tiles were made from eight different manufacturing molds, marked clearly on each tag (fig. 4A). Because 

differences in pre-deployment tile weights were significant 

between molds but negligible within each mold (for a single 
Figure 3. Monitoring and sampling methods. A, Calcification station in the field (note second station in the 

background). B, Slabbing live coral colonies with a modified tile saw. C, Coral colony after it has been sliced. D, Healed 

coral colony, 6 months post-sampling.

6 Methods for Monitoring Corals and Crustose Coralline Algae to Quantify In-Situ Calcification Rates

batch of tiles), it was not necessary to pre-weigh each tile as 

long as the mold number was noted. The whole assembly was 

affixed to the side of the block by sliding two 3-inch hex head 

cap screws through the holes in the nylon rod and the block 

and securing it in place with a flat washer and two machine 

screw hex nuts on each cap screw (fig. 1, table 1). The tile 

was oriented vertically, to more selectively target the CCA 

(as opposed to the fouling) community, and to the side (away 

from the block), to allow grazing organisms equal access to 

both sides of the tile (fig. 3A). The tile was held in place on the 

nylon rod and in the correct orientation by a rubber O-ring and 

4-inch “miniature” cable tie on each side (fig. 1, table 1).

Monitoring and Sampling

Following the initial setup of the calcification stations, 

sites were revisited every 6 months [late spring (May) and late 

fall (November)] to weigh the corals, allowing us to record 

precise changes in mass gained per unit time and detect any 

sub-annual patterns in calcification rates. Corals were only 

stained during springtime visits. Weighing and staining were 

completed in the same manner as in the initial collection, 

except that, during fall (nonstaining) visits, transport buckets 

remained in the boat while corals were weighed on the dock, 

since the corals were out of the ocean for a much shorter 

duration than during springtime (staining) visits. The algal 

A

C

B

C D
Figure 4. Plastic “pear” neck tags, used in the livestock industry, used here as algal recruitment tiles. A, Blank (predeployment) tile, with the manufacturing mold number clearly marked. B, Post-deployment tile, completely encrusted by 

crustose coralline algae after 6 months.

Results and Discussion 7

recruitment tiles also were collected at each visit and replaced 

with new tiles. Harvested tiles were kept in individual, sealable plastic bags and stored on ice or refrigerated (4 °C) until 

they could be processed in the lab. Before reattaching the 

corals and new algal tiles to the blocks, the top of each block 

was cleaned with a rock chisel and brush, and the front portion 

of each nylon rod was scraped clean with a wood chisel. The 

station number plates and coral cap screws also were cleaned 

during each visit. Unless it was the final visit of the monitoring 

period, corals were then returned to the study site and attached 

to new blocks at random.

At the completion of the monitoring period (corals in this 

study were monitored for 2 years), the corals were removed 

from the calcification stations, and two slabs (roughly 3/16 inch, 

or 5 millimeters, thick) were cut from the middle of each colony using a modified wet tile saw in the field, with seawater in 

the cooling bath (figs. 3B, C). To hold the coral in place while 

slicing, the coral was secured to a piece of PVC sheeting by 

inserting the cap screw through a hole drilled in the center and 

fastening it with a wingnut. To position the saw blade for the 

first cut, plastic spacers (approximately 5/16 inch thick) were 

placed between the PVC sheet and the edge of the saw-blade 

guide, and a single spacer was removed after each cut so that 

the thickness of the slabs was as consistent as possible. Using 

a hand-held band saw, the coral was then separated from the 

PVC plate, freeing the two newly cut slabs. The remaining live 

halves of the coral were kept together, and all 10 corals from 

each site were then returned to a location near their collection 

site. Once appropriate locations for the corals were identified, 

the substratum was scraped clean with a rock chisel and brush, 

and the corals were reattached to the seafloor using epoxy. 

Harvested slabs were cleaned with a sonicator in a distilled 

water bath, air dried, and photographed with a stereomicroscope in the lab. The distance from the alizarin red stain line 

to the surface of the coral was measured along the corallite 

wall for each slab, to obtain estimates of linear extension rates, 

using iSolutions® image analysis software.

In preparation for weighing, algal tiles were rinsed 

with fresh water to remove loose sediments (though gently, 

so as not to detach any CCA or invertebrates), photographed 

on both sides, and dried in a 60 °C oven for 48 hours 

(fig. 4B). Once dry, tiles were weighed with and without 

attached invertebrates. Any pieces of CCA that did flake 

off during the rinsing or drying process were included in 

the weight of the tile. The mold-specific mean dry weight 

was then subtracted from the 48-hour (post-deployment) 

dry weight to obtain the mass [grams of calcium carbonate 

(CaCO3)] gained during the 6-month deployment period. 

The number of parrotfish bite marks on each tile was counted, 

using the digital photographs of the tiles, to reveal any patterns 

in grazing pressure.

Results and Discussion

Our calcification station design, in combination with the 

methods employed to measure growth, proved effective for 

monitoring in-situ coral and crustose coralline algal (CCA) 

calcification rates (see Kuffner and others, 2013 for detailed 

results). The station design successfully allowed for the secure 

attachment and periodic removal of coral colonies and algal 

tiles and ensured firm fixation of the blocks to the substratum. 

The majority of the stations have remained intact and in place, 

A B
8 Methods for Monitoring Corals and Crustose Coralline Algae to Quantify In-Situ Calcification Rates

with the exception of a handful of blocks (5 of 40 blocks, in 

the first 4 years of the study) that became loose or, in some 

cases, completely detached from the seafloor as a result of 

strong storms or boats anchoring on the site. Dislodged blocks 

were reinstalled in their original (or, when not possible, a 

nearby) location by drilling and epoxying new rods in place. 

Loose blocks were reinforced using additional epoxy placed 

between the block and the substratum and around the fasteners. In one case, a large piece of reef substratum, which held 

one of the calcification stations, was flipped over during a 

high surf event from Hurricane Sandy, resulting in bleaching 

(caused by lack of light) of the attached coral colony. Corals 

have shown very little mortality while mounted to the calcification stations. To date, only 3 out of 80 colonies have been 

omitted from the dataset: 2 because of severe sponge infestations and 1 because it was removed from the study site by vandals. In fact, new polyps were often visible along the growth 

margins and within the colonies, and most corals grew new 

tissue over the epoxy at the base of the colony, attesting to 

their ability to survive transplantation and their general health 

throughout the monitoring period. Little change was observed 

in the infaunal community (visible from the outside of the 

corals) during the study, indicating that corals were no more or 

less susceptible to infestation while attached to the blocks.

Corals were weighed in November and May to establish 

summer growth rates (measured in November) and winter 

growth rates (measured in May), and rates were subsequently 

compared to elucidate any seasonal patterns in calcification. 

Since we aligned the growth intervals with seasonal boundaries, the weighing intervals used in this study would be applicable worldwide (seasons would simply be reversed in the 

southern hemisphere), enabling the synchronization of calcification-monitoring efforts and the evaluation of spatial patterns 

on a much broader scale. Originally we intended to stain the 

corals during each visit (every 6 months), but, after harvesting 

the first batch of corals in Spring 2011, it was apparent that 

reduced calcification rates during the winter prevented corals 

from incorporating enough dye into their skeleton during 

the 4-hour staining period to produce a distinct band. Corals 

from the Dry Tortugas, where calcification rates were highest 

(Kuffner and others, 2013), were the only ones in which these 

winter stain lines were clearly visible. Thus, we only stained 

our second batch of corals during springtime visits.

Algal recruitment tiles were deployed at every station to 

measure net CCA accretion (it was not possible to measure 

gross accumulation, as tiles were exposed to grazing). Because 

using nylon tags of known weight and surface area removes 

the need for sample dissolution, samples can be archived 

and used for additional analyses (for example, CCA species 

identification for community composition analyses). Only 2 

of the 200 tiles deployed to date have disappeared from the 

stations during their 6-month deployment. The proportional 

contribution of attached invertebrates to the total weight 

accrued on each tile was minimal (mean ± SE = 3.29 plus or 

minus 0.6 percent) and was only noticeable on a small number 

of tiles (82 percent of the tiles had less than 5 percent invertebrate weight), indicating that the algal tiles were successful in 

attracting predominately the CCA community. Invertebrates 

that did settle on the tiles were easily removed for weighing 

(for example, bivalves, fire coral, and worms) or did not add 

a measurable amount of weight to necessitate removal (for 

instance, most sponges and bryozoans). Despite invertebrate 

mass being easy to exclude, competition for tile space would 

potentially be of concern since it has been shown that invertebrates, once settled, have the ability to out-compete slowergrowing CCA for space (Breitburg, 1984). However, instances 

of substantial coverage by these organisms were rare in this 

study; thus, the effect of competition for tile space on CCA 

growth was likely minimal. Spatial differences in CCA growth 

were observed (Kuffner and others, 2013), but the role of 

herbivory and territorial damselfish is unclear and needs to be 

investigated in future studies. Controlling for grazing will be 

necessary to quantify its effect on CCA growth.

Harvest of coral slabs from live colonies in the field, prior 

to reattaching them to the reef, offers a nonlethal method of 

obtaining a complete-colony record of growth, which provides 

an invaluable tool for further studies. Of the sampled colonies 

revisited (30 out of 40), 100 percent survival was attained, 

and the two halves of each coral colony had fused together 

within 6 months after harvesting, leaving only a faint, raised 

seam line where the two halves joined (fig. 3D). We found that 

colonies in which the two halves were not epoxied together 

prior to attachment to the seafloor healed more quickly than 

those that had epoxy between the halves, suggesting that 

direct contact of coral tissue from the two halves promoted the 

healing process. In addition to linear extension measurements, 

numerous other studies can be performed using the coral slabs, 

to further investigate growth parameters and environmental 

variation over the course of skeletal formation. For example, 

corallite morphology (including corallite diameter and spacing) has been shown to vary in response to environmental conditions, such as light intensity or sedimentation rate (Foster, 

1979; Foster, 1980). Such morphological characteristics can 

be assessed within coral slabs to analyze phenotypic patterns 

and attempt to connect them with underlying environmental 

factors. The area of coral skeleton excavated by bio-eroding 

organisms also can be approximated using digital x-rays of 

the slabs, and inferences can be made regarding water quality and water-column productivity based on bio-erosion data 

(Sammarco and Risk, 1990). The relative concentrations of 

strontium and calcium (Sr/Ca) can be measured, using skeletal 

material from the slabs, and used, in conjunction with underwater temperature data, to calibrate this temperature proxy and 

reconstruct historical water temperatures (Beck and others, 

1992) back to the date of coral colony establishment (in this 

case, nearly a decade). Carbonate material from the slabs also 

could potentially be analyzed for the purpose of calibrating 

other trace-element (such as boron and barium) or stableisotope (such as δ18O) proxies for various environmental 

variables.
References Cited 9

Summary

Systematic, long-term monitoring to document current 

spatial and temporal patterns in coral and CCA calcification 

rates will be critical to detecting future changes and assessing 

the effects of ocean warming and acidification on calcification 

and reef-building processes. This study introduces a reliable 

and reproducible method for monitoring calcification rates of 

whole coral colonies and the CCA community. Our goal in 

presenting this guide is to encourage other programs to initiate 

long-term calcification monitoring and to provide specific 

methods to do so, in hopes of improving and synchronizing 

monitoring efforts and enabling the comparison of calcification rates across regions and oceans. A broad, collaborative 

calcification-monitoring effort would be instrumental in 

effectively documenting and quantifying the effects of global 

climate change on coral and algal carbonate deposition and 

reef construction. 

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For further information about this publication contact:

Director

U.S. Geological Survey

St. Petersburg Coastal and Marine Science Center

600 Fourth Street South

St. Petersburg, FL 33701

Or visit the St. Petersburg Coastal and Marine Science Center Web site at:

http://coastal.er.usgs.gov/

Prepared by the Raleigh and the Rolla Publishing Science Center

Morrison and others—Methods for Monitoring Corals and Crustose Coralline Algae to Quantify In-Situ Calcification Rates—Open-File Report 2013–1159