Recruitment of sessile marine invertebrates on ...

BULLETIN OF MARINE SCIENCE, 72(3): 813–839, 2003

RECRUITMENT OF SESSILE MARINE INVERTEBRATES ON HAWAIIAN MACROPHYTES: DO PRE-SETTLEMENT OR POSTSETTLEMENT PROCESSES KEEP PLANTS FREE FROM FOULING? Linda J. Walters, Celia M. Smith and Michael G. Hadfield ABSTRACT Although the polychaete Hydroides elegans and bryozoan Bugula neritina are dominant members of the fouling community in Hawaiian waters, these organisms were rarely found on macroalgae and seagrasses. Three explanations for this lack of fouling on macrophytes were considered: (1) larvae did not settle on these surfaces, (2) physical characteristics of the macrophytes prevented firm attachment, and (3) macrophyte flexibility exceeded animal flexibility. To determine if either invertebrate would settle on 40 species of macrophytes, settlement bioassays were run. When exposure was toxic to larvae, chemical anti-fouling mechanisms were suggested. Macrophytes that were not toxic, but avoided by settling larvae, may have been chemically protected by non-toxic, deterrent compounds, or avoided because of unacceptable morphologies or surface energies. When larval settlement occurred, adhesion strength was tested. To determine if macrophytes were more flexible than attached invertebrates, the macrophyte with attached animals was bent around rods of decreasing diameter until animals were dislodged, or the lamina broke. One or more of the tested types of anti-fouling protection could explain the lack of H. elegans on all 40 macrophytes. Pre-settlement defenses explained the lack of H. elegans on 23 species; post-settlement processes could explain its absence on the remaining 17 species. The lack of B. neritina could only be explained on 18 species of macrophytes.

The negative impact of predators, competitors, and fouling organisms on the fitness of sessile organisms may be reduced if individuals are chemically and physically protected (e.g., Davis et al., 1989; Wahl, 1989; Paul, 1992). A single defense may protect an organism from one biotic threat, but in many habitats multiple defenses are essential to survive the range of interspecific interactions that confront an organism (e.g., Littler et al., 1983; Harvell et al., 1988). Additionally, there may be additive or synergistic effects when defenses are combined (e.g., Gerhart et al., 1988; Hay et al., 1994). Prey in marine habitats may be protected chemically by secondary metabolites and morphologically by secreting hard spines or spicules (e.g., Littler and Littler, 1980; Harvell, 1984; Lindquist et al., 1992; Pennings and Paul, 1992; Van Alstyne and Paul, 1992). Chemical and physical defenses can also influence the outcome of competitive interactions in corals, anemones, sponges and macroalgae (e.g., Sammarco et al., 1983; Bak and Borsboom, 1984; Porter and Targett, 1988). For example, contact with secondary metabolites on the surface of the red alga Plocamium hamatum caused tissue necrosis in the soft coral Sinularia cruciata (de Nys et al., 1991). Physical defenses that can influence the outcome in a competitive situation include nematocyst-bearing structures, mucus, extra-coelentaric digestion, spine production, and thickening at contact margins (e.g., Lang, 1973; Chadwick, 1988; Buss, 1990; Keats and Maneveldt, 1994). Like predation and competition, macrofouling can significantly reduce the fitness of the host (see reviews by Davis et al., 1989; Wahl, 1989). Surfaces of macroalgae and seagrasses provide an important spatial resource for the attachment and growth of sessile marine organisms. However, epiphytes can reduce photosynthetic capacities and increase drag on hosts, and this frequently results in whole or partial plant mortality, reduced 813

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BULLETIN OF MARINE SCIENCE, VOL. 72, NO. 3, 2003

growth rates, and decreased reproductive output (e.g., Crisp and Williams, 1960; Woollacott and North, 1971; Bernstein and Jung, 1979; but see Cancino et al., 1987; Muñoz et al., 1991). Additionally, fouling organisms may increase levels of host herbivory, as is the case with the epiphytic red alga Ceramium codicola on its host, the green alga Codium fragile (Trowbridge 1993). To date, many studies of the chemicals that reduce fouling of marine macrophytes have focused on chemical extracts of macerated algal or seagrass tissues (e.g., Sieburth and Conover, 1965; Todd et al., 1993). For chemical protection to occur, however, compounds must either be found on the exposed surfaces of the plants, or are released into the water column (Walters et al., 1996). Toxic compounds may kill competent larvae, while sublethal concentrations of these compounds may poison recruits over time. Walters et al. (1996) documented that water conditioned with the brown alga Dictyota crenulata (published as D. sandvicensis) killed all larvae of the serpulid polychaete Hydroides elegans and the bryozoan Bugula neritina within minutes of exposure in laboratory assays. Alternatively, if a competent larva encounters a non-toxic, inhibitory compound, settlement may occur elsewhere. Schmitt et al. (1995) found that larvae of Bugula neritina rarely settled on Dictyota menstrualis, even though in laboratory choice trials it contacted the surface of this plant as frequently as the preferred algal host Gracilaria tikvahiae. If secondary chemicals do not prevent recruitment of sessile invertebrates on marine plants, then physical characters may deter larval settlement, or remove attached individuals. Plant attributes that may influence settlement by fouling species may be related to topographic complexity, overall plant dimensions, surface free energy (wettability), or mucus secretions (e.g., Crisp and Ryland, 1960; Williams 1964; Mihm et al., 1981; Rittschof et al., 1988; Walters and Wethey, 1991). Once attached to a plant surface, epiphytic organisms may be removed by regular sloughing of the epidermis, or portions of the outer cell walls (e.g., Filion-Myklebust and Norton, 1981; Russell and Veltkamp, 1984; Johnson and Mann, 1986). The existence of additional post-settlement plant defenses has been inferred (Seed and O’Connor, 1981). For example, if surface characteristics of a thallus prevent firm attachment, then the chances of dislodgment increase as water motion increases and the size of the epiphyte increases. A second possible postsettlement mechanism that may remove epiphytes involves plant flexibility. If the plant is more flexible than the epiphyte, then attached epiphytes may be dislodged when water motion results in plant flexure exceeding epiphyte flexibility. Many have postulated that combined chemical and physical pre- and post-settlement anti-fouling defenses keep marine plant surfaces clean (e.g., Bakus et al., 1986; Davis et al., 1989; Wahl, 1989; Paul, 1992; Pawlik, 1992). Here, we address this possibility with a diverse assemblage of tropical marine plants collected from shallow waters around the island of Oahu, Hawaii. Although sessile invertebrates are commonly found on the surfaces of macroalgae and seagrasses in temperate waters (see reviews by Crisp, 1976, 1984; Seed, 1986), many long-lived, intertidal and shallow subtidal Hawaiian macroalgae and angiosperms are rarely fouled by sessile invertebrates (Magruder and Hunt, 1979; Michael and Smith, 1995). The mechanisms that enable these plants to remain clean are the focus of this study. First, we quantified the extent of fouling by sessile invertebrates on 40 common marine plant species in the field. For all species of which the number of invertebrate recruits was small, we considered three potential explanations for the lack of fouling: (1) larvae avoid settling on plant surfaces, (2) plants have surface characteristics that deter long-term attachment, and (3) blade flexibility exceeds that of rigidly attached

WALTERS ET AL.: FOULING PREVENTION BY MARINE PLANTS

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invertebrates. If none of these hypotheses are accepted, then the potential exists that the tested animals readily settle on marine plants, but are regularly removed by other processes, such as predation, or that larvae of these animals rarely encounter these plants in the field. METHODS QUANTIFICATION OF FOULING ON HAWAIIAN MARINE PLANTS.—The extent of fouling by sessile invertebrates was determined for 40 species of marine plants that represent common members of the Phaeophyta, Chlorophyta, Rhodophyta, Cyanobacteria and seagrasses around the island of Oahu (Table 1). All are found intertidally, or in shallow reef areas (≤ 4 m) in mixed-age assemblages. Fifty intact, healthy adult plants of each species were collected from three of the eight possible field collection locations (Fig. 1) on three dates between February 1992–December 1993 (Table 1). Collection locations were determined by macrophyte availability. For species found only at one field location, 50 intact plants were collected from the site in three different months. Collection dates for all other species differed for each macrophyte, based on seasonal differences in mature macrophyte availability. In total, 150 individuals of each species were censused during the course of this study. This collection strategy was employed to maximize the potential for detecting differences among sites, while minimizing the number of harvested individuals. Plants were kept in seawater-filled, five-gallon buckets until arriving at the Kewalo Marine Laboratory in Honolulu (< 1 hr), where they were immediately placed in shallow, outdoor aquaria supplied with running-seawater. To detect and quantify the presence of sessile invertebrates on each plant species, all individuals were examined at 10× with a dissecting microscope, and the presence of all epiphytes was recorded. Only plants that were free from fouling by invertebrates and epiphytic algae were used in the laboratory trials. SESSILE INVERTEBRATES USED IN LABORATORY TRIALS.—The tube-building serpulid polychaete H. elegans and the arborescent bryozoan B. neritina were used in all experiments. Both were introduced to Oahu, and established populations of both can now be found island-wide (Eldredge and DeFelice, 2000). These species were chosen for this study because they are two of the dominant fouling organisms in Hawaii, and large numbers of competent larvae may be readily obtained from both (Edmondson, 1944). Hydroides elegans were found at all eight collection sites (Fig. 1), predominately on artificial or dead substrata (coral rubble, shells, pilings, boat hulls, etc.). Bugula neritina is found attached to marine plants in many temperate and tropical habitats around the globe (e.g., Keough, 1986; Walters and Wethey, 1991). However, in Hawaii individuals were found attached almost exclusively to man-made substrata in harbors. At the locations where plants were collected, only a few individuals of B. neritina were observed on shells and rubble. Adults of H. elegans were collected from Hospital Point, Pearl Harbor, allowed to spawn in the laboratory, and their larvae reared to metamorphic competence within six days using methods described in Hadfield et al. (1994). Colonies of B. neritina release larvae that are competent to settle (Lynch, 1947). Adults were collected from either Rainbow Marina (Pearl Harbor) or Ala Wai Marina (Honolulu), and kept in running-water sea tables covered with black plastic until larvae were required. Competent larvae could then be collected by exposing adults to light (Lynch, 1947). Behaviors of larvae of B. neritina collected this way are similar to those of field-collected larvae (L. J. Walters, per. obs.). LARVAL SETTLEMENT ON PLANTS IN THE LABORATORY.—From the field observations, it was not possible to determine if larvae (1) never contacted plants, (2) contacted surfaces but did not settle on them, or (3) attached to the plants and were removed before our observations. To ensure larvae contacted plants and to determine the extent of settlement on plant surfaces under controlled conditions, we ran separate laboratory bioassays with plants from the three collection locations (or three dates from one collection location) and larvae of H. elegans and B. neritina (Table 1). In each trial, three replicate dishes were tested for each plant species. Particulate matter was removed from each


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Table 1. Survival and settlement location preferences of larvae of Hydroides elegans and Bugula neritina. Collection sites correspond to locations in Figure 1. The sequence of dates corresponds to the sequence of collection sites for both H. elegans and B. neritina. Additionally, the dates for the H. elegans trials correspond to the dates when evidence of fouling was examined on 50 replicates of each macrophyte. Survival values are followed by a * for plant species with significantly reduced survival in settlement bioassays when all plant species were compared simultaneously with ANOVA (H. elegans: P < 0.05; B. neritina: : P < 0.05; N = 3) and Tukey multiple comparisons tests. For plant species that were not toxic to larvae, Preference = location where larvae preferentially settled in dishes when compared with ANOVA (H. elegans: : P < 0.05; B. neritina: : P < 0.05; N = 3) and Tukey tests after normalizing by available surface area. Hydroides elegans Chlorophyta Avrainvillea amadelpha Bornetella sphaerica Bryopsis pennata Caulerpa racemosa Caulerpa taxifolia Caulerpa verticillata Cladophora dotyana Codium arabicum Codium edule Codium reediae Dictyosphaeria cavernosa Enteromorpha flexuosa Halimeda discoidea Microdictyon setchellianum Neomeris annulata Ulva fasciata Ulva reticulata Valonia aegagropila Ventricaria ventricosa Phaeophyta Colpomenia sinuosa Dictyota acutiloba Dictyota crenulata Lobophora variegata Padina australis Sargassum echinocarpum Sargassum polyphyllum Sphacelaria tribuloides Turbinaria ornata Rhodophyta Acanthophora spicifera Asparagopsis taxiformis Galaxaura rugosa Gracilaria coronopifolia Hypnea musciformis Jania capillacea Kappaphycus alvarezii Laurencia cartilaginea Liagora tetrasporifera Melanamansia glomerata Cyanobacteria Lyngbya majuscula Angiosperma Halophila hawaiiana Control polystyrene

Site(s)

Dates

Survival (SE)

3 1, 3, 5 1, 2, 6 1, 2, 6 6 6 3, 4, 7 1, 6, 7 2, 3, 7 1, 2, 6 1, 3, 6 1, 2, 5 1, 2, 3 1, 3, 4 1, 3, 4 4, 7, 8 1 1 6

11/92, 12/92, 2/93 2/92, 11/92, 10/93 2/92, 3/93, 10/93 2/92, 4/92, 11/92 2/93, 4/93, 5/93 10/92, 2/93, 3/93 11/92, 6/92, 5/92 2/92, 11/92, 1/93 3/92, 1/93, 1/93 3/92, 3/93, 2/93 2/92, 1/93, 3/93 2/93, 4/92, 10/93 2/92, 3/92, 11/92 2/92, 3/93, 6/92 3/92, 11/92, 6/92 6/92, 5/92, 4/93 2/92, 2/93, 4/93 2/92, 2/93, 4/93 10/92, 4/93, 5/93

100.0 (0) 100.0 (0) 100.0 (0) 100.0 (0) 100.0 (0) 100.0 (0) 56.4 (14.1)* 100.0 (0) 99.0 (0.9) 100.0 (0) 100.0 (0) 100.0 (0) 21.3 (5.9)* 100.0 (0) 38.7 (21.0)* 100.0 (0) 100.0 (0) 88.75 (15.9) 100.0 (0)

1, 2, 7 1, 3, 5 1, 4, 6 1, 3, 7 1, 4, 7 2, 4, 7 1, 4, 7 1, 6, 7 1, 2, 4

2/92, 4/92, 1/93 10/93, 10/93, 3/93 2/92, 6/92, 11/92 3/92, 11/92, 1/93 2/92, 6/92, 5/92 3/93, 6/92, 5/92 2/92, 6/92, 5/92 2/93, 3/93, 3/93 2/92, 3/92, 6/92

69.9 (13.5) 100.0 (0) 56.6 (19.0)* 68.86 (14.3) 85.4 (16.9) 100.0 (0) 100.0 (0) 100.0 (0) 78.9 (22.1)

1, 2, 4 1, 2, 3 2, 3, 4 1, 2, 7 1, 2, 6 3, 4, 7 6 1, 2, 7 2, 3, 7 1, 3, 8

2/92, 3/92, 6/92 2/93, 3/93, 3/93 3/93, 1/93, 6/92 2/92, 4/92, 5/92 2/93, 4/92, 10/92 3/93, 6/92, 5/92 10/92, 2/93, 3/93 2/93, 3/92, 1/93 4/92, 11/92, 5/92 2/92, 11/92, 3/93

100.0 (0) 52.8 (21.6) 98.1 (2.3) 79.8 (23.3) 100.0 (0) 83.8 (19.9) 59.4 (19.6)* 98.6 (2.3) 45.0 (17.2)* 62.5 (16.3)

1, 3, 6

3/92, 11/92, 11/92

1, 3, 7

2/92, 3/93, 5/92

Bugula neritina Dates

Survival (SE)

Preference

3/93, 7/93, 8/93 3/92, 3/93, 4/93 2/92, 3/92, 7/93 2/92, 4/92, 7/93 4/93, 5/93, 6/93 7/93, 8/93, 9/93 3/93, 7/93, 7/93 2/92, 7/93, 7/93 3/92, 3/93, 7/93 7/93, 3/93, 7/93 2/92, 3/93, 7/93 7/93, 4/92, 4/93 2/92, 3/92, 3/93 2/92, 3/93, 7/93 3/92, 3/93, 7/93 7/93, 6/92, 4/93 2/92, 4/93, 7/93 2/92, 4/93, 5/93 3/93, 7/93, 8/93

100.0 (0) 100.0 (0) 99.4 (0.4) 100.0 (0) 100.0 (0) 99.9 (0.1) 100.0 (0) 98.8 (0.5) 100.0 (0) 100.0 (0) 99.6 (0.4) 99.8 (0.2) 99.9 (0.1) 100.0 (0) 99.2 (0.8) 100.0 (0) 100.0 (0) 100.0 (0) 100.0 (0)

dish pant

2/92, 4/92, 7/93 10/93, 10/93, 10/93 2/92, 7/93, 7/93 3/92, 3/93, 7/93 2/92, 7/93, 7/93 3/93, 7/93, 7/93 2/92, 7/93, 6/92 3/93, 8/93, 7/93 2/92, 3/92, 7/93

87.5 (7.6)* 100.0 (0) 100.0 (0) 100.0 (0) 99.9 (0.1) 99.7 (0.3) 100.0 (0) 87.4 (6.3)* 100.0 (0)

2/92, 3/92, 7/93 2/92, 4/93, 3/93 3/93, 3/93, 6/92 2/92, 4/92, 7/93 4/93, 4/92, 7/93 3/93, 7/93, 6/92 7/93, 8/93, 9/93 3/92, 3/92, 7/93 4/92, 3/93, 5/92 10/93, 3/93, 3/93

100.0 (0) 90.6 (4.9) 98.1 (1.1) 100.0 (0) 100.0 (0) 90.1 (5.6)* 100.0 (0) 97.4 (1.9) 99.2 (0.3) 99.4 (0.6)

52.1 (21.7)*

3/92, 3/93, 7/93

73.8 (8.1)*

100.0 (0)

2/92, 7/93, 6/92

100.0 (0)

100.0 (0)

Preference

dish dish dish

dish dish

dish dish plant

dish dish dish dish

dish dish dish dish plant

dish

dish plant dish dish plant

plant

dish

dish plant dish

plant

100.0 (0)

plant surface, and then, depending on plant dimensions, small, haphazardly selected pieces of plant biomass from different individuals or entire plants were placed in separate, sterile, 60 × 15 mm, polystyrene petri dishes to provide a similar plant surface area (approximately 25 × 20 mm) in each dish. Then, 15 ml of 0.45 µm-filtered seawater was added to each dish. Three control dishes contained only filtered seawater. All dishes were kept overnight under similar laboratory conditions (ambient seawater temperature: 23°C; overhead fluorescent lighting: 14 h L, 10 h D; irradiance ≈ 15 µmol m−2 s−1). After 16 h, the water was changed and the larval settlement trials begun. This acclimation period was to allow any chemical or biological materials that were the result of damage during collection or experimental preparation to be leached from the plants. Unlike many temperate macrophytes that quickly decay with such handling, many tropical plant species rapidly heal (Walters and Smith, 1994; Smith and Walters, 1999). In preliminary trials, plants that survived this time period recovered from any wounds, did not obviously continue to leach biological or chemical


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Figure 1. Plant collection locations on Oahu, Hawaii, and the mean maximum flow rates in cm s−1 (± S.E.) at each location during the most dynamic coastal weather conditions in 1993 (December 18–30). materials, and remained healthy and continued to grow in the laboratory for a minimum of one month (L. J. Walters, per. obs.). Trials with H. elegans and B. neritina were run simultaneously to the extent made possible by plant and larval availability (Table 1). Hydroides elegans was present in the field throughout the 23 mo period; B. neritina was available from February 1992–June 1992 and from March–December 1993. The bioassay protocol followed Walters et al. (1996). Briefly, for the settlement trials with H. elegans, approximately 100 larvae, 0.45 ml of a culture of the phytoplankton Isochrysis galbana (Tahitian strain; supplied at a concentration of ca. 6 × 105 cells ml−1), and a 28 × 21 mm piece of black plastic mesh (Vexar, mesh size: 7 × 6 mm) that had been soaked for three d in running

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seawater to develop a biological film were added to each test or control dish. The biofilmed mesh was added to provide larvae with a positive settlement cue (Hadfield et al., 1994; Unabia and Hadfield, 1999). New batches of larvae were used in each trial, and each batch was produced from gametes of at least ten adults. Dishes were placed in a haphazard array on a laboratory bench under the light and temperature conditions described above. Additional I. galbana were added to each dish every other day throughout the trial. With B. neritina, after 100 larvae were added to each dish, the dish was immediately covered with aluminum foil to prevent light from influencing settlement of these photonegative larvae (Lynch, 1947). This technique is commonly used for settlement assays in which the test larvae respond to light cues (e.g., Walters et al., 1996). Plastic mesh was not added to these dishes because larvae of B. neritina do not require a biofilm cue for settlement (Walters et al., 1996). Trials ended when all larvae in control dishes had settled (5–7 d for H. elegans; 24 h for B. neritina). Survival of larvae in all dishes was then calculated. Data were arcsine transformed so that statistical analyses could be run on normally distributed data with homogeneous variances (Sokal and Rohlf, 1995). Mean survival of animals in dishes for each plant species was then compared simultaneously with a two-way nested analysis of variance (ANOVA) with plant species and collection location as the main effects (SAS 6.08, SAS Institute, 1988). Tukey a posteriori multiple comparisons tests were run to determine differences between treatments. No further analyses were run for plant species if larval survival was significantly reduced (ANOVA, P < 0.05). For all other plants, the location of all settled individuals in each dish was recorded as on the: (1) plant surface, (2) plastic mesh (H. elegans), or (3) submerged sides and bottom of the petri dish. During the course of the trials with H. elegans, a biological film developed on the submerged dish surfaces, and larvae settled on this substratum as well as on the mesh. Thus, the areas of the mesh and the dish were combined for the analyses. The total surface area available for settlement on each plant, the mesh, and the dishes was calculated using an image-analysis system (Mocha Version 1.2.10, Jandel Scientific, Inc.). If settlement was random, there should have been the same number of settlers/mm2 on the alga as on the dish (B. neritina) or on the dish + mesh (H. elegans). The difference between the number of settlers/mm2 observed on each alga and the number of settlers/mm2 on the dish (or dish + mesh) were compared simultaneously for all non-toxic plant species at all tested locations using a two-way ANOVA (main effects: plant species and collection location; N = 3) and Tukey multiple comparisons tests. POST-SETTLEMENT REMOVAL OF SESSILE INVERTEBRATES: ADHESION TO PLANT SURFACES.—To understand the importance of water motion on the adhesion strength of sessile invertebrates to marine plants, we determined the maximum flow rates experienced by plants in the field and then subjected attached animals to flow rates up to this maximum under controlled conditions. Maximum flow rates in the field were calculated using maximum-velocity flow recorders designed after those described by Bell and Denny (1994). Our recorders accurately measured maximum flow rates between 10–50 cm s−1. Six recorders were placed in representative locations in each of the eight collection locations for one tidal cycle. Recorders were attached in the field with swivels and cable ties where marine plants were common and the devices would not become entangled. Maximum flow was recorded during the most dynamic coastal weather conditions of 1993 (December 18–30: 20–30 mph winds, 5–10 ft waves). No post-settlement trials were run with plant species that were (1) toxic or (2) non-toxic and avoided by 100% of settling larvae of H. elegans or B. neritina. For all others, larvae were allowed to settle and grow on plant surfaces in still water in the laboratory for two d (new recruits), seven d (juveniles), or until the animals were reproductively mature (30 d H. elegans; 15 d B. neritina). The dimensions of each age-class animals grown under these controlled conditions were similar (ANOVA, P > 0.05). On alternate days, the water was changed in each dish, and an excess amount of the phytoplankton I. galbana was added as food. When animals reached the specified age, individuals were subjected to a stream of water at pre-determined speeds until the animal was removed, or subjected to the maximum water pressure.


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The water-jet device consisted of a 20 L Nalgene carboy with a 16 mm diameter spigot. The carboy was placed on the second floor balcony of the laboratory. From the spigot, a length of 18 mm diameter plastic tubing hung straight down 4.8 m. With additional tubing, the inner diameter was reduced from the spigot width to 5 mm. A water-pick tip (Teledyne Inc.) was attached to the free end of the tubing (inner diameter of the open end of the water-pick tip: 1 mm; tip length: 6.5 cm). To keep the head pressure constant, the carboy was always overflowing. A constant, reproducible flow of seawater was created by placing a series of clamps on the tubing. Specific flow rates used in our trials were 10, 21, 32, 43, and 54 cm s−1. Macrophytes were held in place with forceps in still water in a glass bowl and observed with a dissecting microscope while individuals of H. elegans and B. neritina on their surfaces were subjected to incrementally greater flow rates with the water-jet. Although this methodology creates a situation where temporal pseudoreplication occurs, we suggest that this mimics reality because water flow in the field varies continually and individuals are exposed to this variation throughout their lifetimes. Additionally, in pilot experiments we found no differences in the results when individuals were subjected to one flow rate versus increasing flows. The water-pick tip was held 1 mm from each test animal (bryozoan or polychaete). Individuals were subjected to each flow rate for one min. For H. elegans, water was directed toward the tube opening for 30 s, and then toward the midpoint along the length of the tube for 30 s. For B. neritina, water at each flow rate was directed at the lophophore of the attachment zooid for 30 s, and then 90º from the front of this zooid for 30 s. Trials were run for each animal-age category (new recruits, juveniles, mature individuals) on each macrophyte host until a minimum of 30 individuals of each age and all attached individuals from a minimum of three plants of each species from each collection location used in the settlement assays were tested. The total number of animals of each age tested for a plant species ranged from 30–185 on a minimum of nine plant surfaces (maximum of 30 surfaces). We acknowledge that having more than one animal per plant host replicate (in some cases) violates the assumption of independence of data; we used this protocol when we could not locate 90 individuals of a plant species (30 for each animal-age category) on a collecting trip. As a conservative estimate of plant protection from fouling, the flow rates required to remove 90% of the H. elegans and B. neritina from each plant were determined. Flow rates were compared for the three age classes of H. elegans and B. neritina. These flow rates were also compared to the flow rates required to remove H. elegans and B. neritina grown under similar conditions on polystyrene control surfaces. We do not present mean flow rates for removal because that precludes inclusion of individuals that remained attached to a plant surface at all flow rates. Additionally, due to the above-mentioned problems with pseudoreplication and independence, only summary statistics are presented. POST-SETTLEMENT REMOVAL OF SESSILE INVERTEBRATES: FLEXIBILITY OF MACROPHYTES.—If plants were more flexible than attached invertebrates, then the latter should be shed or damaged when plants bent in response to water motion. Plants with attached 7-d H. elegans or B. neritina were bent around rods of decreasing outer diameter (15.5, 14.0, 12.5, 11.0, 9.5, 8.0, 6.5, 5.0, 3.5, 2.0, 1.0, 0.5 mm) in still water at a constant rate (1 mm s−1). If the tissues of a plant did not break within this range, then its surfaces were bent together until the plant tissues broke or all surfaces were in contact and flexure was 360º (not breakable). Additionally, the rod diameters at which animals were removed from plant surfaces (intact or in pieces) were recorded. As with the adhesion trials, pseudoreplication exists with this protocol. However, this methodology likely mimics natural conditions with continual changes in flow rates. All 7-d individuals used in these trials were attached to different plants, including a minimum of three plants from each collection location reported in Table 1 (N = 30). From these data, we calculated the radius of curvature (Cr) at which plant breakage and animal removal occurred. In our trials, Cr was equal to 0.5 (circumference of the rod; Walters and Wethey, 1991). By comparing the Cr at which tissues from each plant broke and the value at which 90% of the attached animals were removed, we determined the potential for each animal species to adhere to plant surfaces under field conditions where plants may be bent to their fullest. As above, 90%

820


removal was selected as a conservative estimate to better understand the importance of this type of post-settlement mechanism for removing fouling organisms.

RESULTS FIELD SETTLEMENT PATTERNS.—Hydroides elegans were present only on one plant species in the field; three H. elegans were found on two individuals of the green alga Dictyosphaeria cavernosa collected from Coconut Island in March 1993. Bugula neritina was not found on any of the 6000 plants examined in any month during the 23 mo study. Additionally, no other sessile invertebrates were found on any plant surfaces, although numerous plants had coralline or filamentous algae covering portions of their surfaces and clonal ascidians and sponges had been previously observed on the inner, protected surfaces of D. cavernosa (M.G. Hadfield, per. obs.). LABORATORY SETTLEMENT TRIALS.—Because fouling by sessile invertebrates was extremely limited on field-collected plants, all 40 species were used in laboratory settlement trials. Mean survival and settlement location preference are given separately for H. elegans and B. neritina in Table 1. For H. elegans, significantly greater mortality than in control dishes (ANOVA; P < 0.05) was associated with proximity to eight macrophytes (Table 1). 15 additional plant species were avoided by settling larvae of H. elegans (Table 1). Larvae of H. elegans rarely settled on plant surfaces more often than would be expected by chance (Table 1). Preferential settlement on plant surfaces was found only in two species of algae: the green alga Valonia aegagropila and the red alga Laurencia cartilaginea. Significant mortality of larvae of B. neritina (ANOVA; P < 0.05) was observed with the cyanobacteria Lyngbya majuscula, two species of brown algae, Colpomenia sinuosa and Sphacelaria tribuloides, and two species of red algae, Asparagopsis taxiformis and Jania capillacea (Table 1). Non-toxic algae that were avoided by settling larvae of B. neritina (ANOVA; P < 0.0001) included eight additional plant species (Table 1). Larvae of B. neritina preferentially settled on Bornetella sphaerica, Microdictyon setchellianum, Ventricaria ventricosa, Sargassum polyphyllum, Laurencia cartilaginea, and Halophila hawaiiana (Table 1). Mean maximum flow ranged from 17 to >50 cm s−1 at our collection sites and is presented separately for each site in Figure 1. The values for the Hale’iwa site were based on five recorders, as the sixth was damaged. Additionally, at the location where water motion was greatest, Makapu’u Beach Park, the mean maximum flow rate could only be inferred as one recorder reached the maximum recording value (50 cm s−1), and the remaining five recorders were either damaged or lost during the 12 h sampling period (the 30 lb test monofilament broke on three flow recorders; the 60 lb strength cable ties failed on the two lost recorders). For H. elegans, collection location significantly influenced larval settlement and attachment location (ANOVA; P < 0.05). Survival in dishes with plants collected at Makapu’u Beach Park (maximum flow rate: > 50 cm s−1) was significantly greater than survival on plants collected at much lower maximum flows (Kaimana Beach: 16.7 cm s−1, Kahala: 19.9 cm s−1, Malaekahana Beach Park: 28.8 cm s−1; Fig. 1, Table 1). However, in laboratory settlement trials, larvae of H. elegans settled in greater numbers on plants from the three lowest flow locations than on plants from the sites with the two highest maximum flow rates (Fig. 1). No consistent patterns relative to flow rate were found for enhanced


821

survival (Malaekahana Beach Park > Ka’alawai Beach and Coconut Island) or settlement location (Kailua Beach Park > Malaekahana Beach Park and Makapu’u Beach Park) with larvae of B. neritina (Fig. 1, Table 1). POST-SETTLEMENT SUCCESS: ADHESION TO MACROPHYTE SURFACES.—The flow rates required to remove 90% of the individuals of H. elegans from plant surfaces generally spanned the range of flows available with our water-jet (Table 2). On control polystyrene surfaces, only the highest flow rate (54 cm s−1) removed 90% of the newly recruited (2-d) H. elegans from the substrata. No juvenile or mature (7-d or 30-d) individuals were removed from polystyrene at this flow rate (Table 2). In contrast, 2-d and 7-d individuals of H. elegans were not as securely attached to any plant surface, and all were dislodged at flow rates of ≤ 43 cm s−1 (Table 2). However, more than 10% of 30-d H. elegans remained attached to eight of the 29 tested plant species when subjected to 54 cm s−1 flows (Table 2). Several additional patterns emerge in examining these results (Fig. 2, Table 2). On some plants, the flow rate required to dislodge H. elegans remained uniform and low over time (10 cm s−1: Codium arabicum, C. edule, C. reediae, Sargassum echinocarpum; 21 cms−1: Caulerpa racemosa; Fig. 2, Table 2). In contrast, on surfaces of Enteromorpha flexuosa, individuals of H. elegans of all ages required substantial force (43 cm s−1) to be dislodged (Fig. 2, Table 2). With many other macrophyte species, the flow rates necessary to remove H. elegans changed as the animals aged. The flow rates required to dislodge individuals increased on some macrophytes (e.g., Hypnea musciformis), and decreased on others (Dictyosphaeria cavernosa; Fig. 2, Table 2). In a few trials, the flow rate necessary to remove recruits of H. elegans increased and then decreased (Colpomenia sinuosa), or following a decrease in the flow rates to remove individuals from day 2–day 7, flow had to be increased to remove adults (from day 7–day 30) (Ulva fasciata, Gracilaria coronopifolia; Fig. 2). Additionally, when individuals of H. elegans of any age were removed from a surface at a flow rate of ≤ 21 cm s−1, they were always removed intact. At higher flow rates, tubes were frequently broken before being removed. All individuals of all ages of B. neritina remained attached to the control surfaces at the maximum tested flow rate (54 cm s−1) in our trials (Table 2). Additionally, it was not possible to remove 90% of the individuals of B. neritina from one or more tested agegroups at the highest flow rate for 23 of the 32 plant species tested (Table 2). When removed from a plant surface, individuals of B. neritina were always intact. As with H. elegans, over time the flow rate required to remove 90% of the individuals of B. neritina on some plants increased (e.g., Halophila hawaiiana), decreased (e.g., Codium reediae), or remained constant (32 cm s−1: Lobophora variegata, Padina australis; Table 2). POST-SETTLEMENT SUCCESS: PLANT VS ANIMAL FLEXIBILITY.—The values for radii of curvature (Cr) at which plant tissues broke or 90% of the animals were removed (intact or in pieces) reveal much greater flexibility among the plants than the animals (Fig. 3, Table 3). Individuals of H. elegans were removed intact from some surfaces, while on other surfaces the tubes were broken (Fig. 3, Table 3). Tube fragments remained attached to a few plants (Enteromorpha flexuosa, Ulva fasciata, S. polyphyllum), but came off most (Fig. 3, Table 3). If B. neritina was dislodged from a plant surface, it was always removed intact (Table 3). Some plant species were extremely flexible and were not broken using this protocol (e.g., Ulva reticulata, Dictyota acutiloba, and Lyngbya majuscula), while others were totally inflexible (Bornetella sphaerica, Dictyosphaeria cavernosa, Ventricaria ventricosa; Fig. 3, Table 3). In two cases, fragments of H. elegans were removed at the same Cr at


822

Table 2. Attachment strength of Hydroides elegans and Bugula neritina to macrophytes. For new recruits (2−d), juveniles (7−d), and adults (30−d H. elegans; 15−d B. neritina), the water velocity required to remove 90% of the individuals on each plant species up to a maximum velocity of 54 cm s−1 is presented (N ≥ 30). No Set = survival was significantly reduced or no larvae settled on the plant species, so no adhesion trials could be run. Change = change in flow rate required to remove individuals from day 2−day 7, and from day 7−day 15 (B. neritina) or day 30 (H. elegans). ↑ represents an increase in flow rate to remove 90% of the individuals as animals aged, ↓ represents a decrease over time, = represents no change over time.

2 d (N)

Hydroides elegans 7 d (N) 30 d (N)

Change

2 d (N)

Bugula neritina 7 d (N) 15 d (N)

Change

Chlorophyta Avrainvillea amadelpha

21.0 (47)

32.0 (185)

32.0 (82)

↑ ,=

>54.0 (30)

>54.0 (58)

32.0 (135)

=,↓

Bornetella sphaerica

10.0 (40)

21.0 (31)

32.0 (71)

↑ ,↑ No Set

21.0 (40)

32.0 (31)

32.0 (71)

↑ ,=

Bryopsis pennata

No Set

No Set

No Set

>54.0 (30)

>54.0 (84)

>54.0 (34)

=,=

Caulerpa racemosa

21.0 (46)

21.0 (52

21.0 (170)

=,=

>54.0 (50)

32.0 (32)

>54.0 (43)

No Set

↓ ,↑ No Set

Caulerpa taxifolia

10.0 (87)

21.0 (51)

21.0 (50)

↑ ,=

No Set

No Set

Caulerpa verticillata

21.0 (51)

21.0 (51)

54.0 (44)

>54.0 (31)

>54.0 (44)

>54.0 (40)

=,=

Cladophora dotyana

No Set

No Set

No Set

=,↑ No Set

54.0 (47)

>54.0 (37)

>54.0 (41)

↑ ,=

Codium arabicum

10.0 (164)

10.0 (30)

10.0 (68)

=,=

10.0 (32)

21.0 (35)

21.0 (68)

↑ ,=

Codium edule

10.0 (115)

10.0 (57)

10.0 (107)

=,=

21.0 (51)

10.0 (37)

21.0 (35)

↓ ,↑

Codium reediae

10.0 (65)

10.0 (62)

10.0 (36)

=,=

21.0 (77)

21.0 (45)

10.0 (63)

=,↓

Dictyosphaeria cavernosa

32.0 (33)

21.0 (37)

10.0 (133)

↓ ,↓

10.0 (62)

>54.0 (44

21.0 (114)

↑ ,↓

=,= No Set

>54.0 (48)

>54.0 (55)

>54.0 (46)

=,=

43.0 (49)

>54.0 (45)

54.0 (67)

↑ ,↓

↑ ,= No Set

43.0 (54)

>54.0 (34)

21.0 (34)

↑ ,↓

43.0 (42)

>54.0 (33)

>54.0 (32)

↑ ,= =,=

Enteromorpha flexuosa

43.0 (35)

43.0 (39)

43.0 (97)

Halimeda discoidea

No Set

No Set

No Set

Microdictyon setchellianum

10.0 (34)

21.0 (40)

21.0 (52)

Neomeris annulata

No Set

No Set

No Set

Ulva fasciata

32.0 (54)

21.0 (30)

>54.0 (49)

↓ ,↑

>54.0 (48)

>54.0 (84)

>54.0 (77)

Ulva reticulata

10.0 (53)

32.0 (82)

>54.0 (102)

↑ ,↑

>54.0 (34)

>54.0 (61)

>54.0 (58)

Valonia aegagropila

10.0 (101)

21.0 (37)

32.0 (81)

↑ ,↑

>54.0 (49)

21.0 (102)

32.0 (40)

↓ ,↑

Ventricaria ventricosa

10.0 (31)

10.0 (49)

21.0 (50)

=,↑

32.0 (79)

54.0 (83)

32.0 (71)

↑ ,↓

Phaeophyta Colpomenia sinuosa

10.0 (39)

21.0 (70)

10.0 (70)

↑ ,↓

No Set

No Set

No Set

Dictyota acutiloba

32.0 (34)

43.0 (44)

>54.0 (39)

>54.0 (58)

>54.0 (36)

>54.0 (31)

=,=

Dictyota crenulata

No Set

No Set

No Set

>54.0 (58)

>54.0 (36)

>54.0 (31)

=,=

Lobophora variegata

10.0 (36)

10.0 (37)

21.0 (69)

=,↑

32.0 (31)

32.0 (34)

32.0 (55)

=,=

Padina australis

21.0 (84)

32.0 (56)

>54.0 (43)

↑ ,↑

32.0 (110)

32.0 (168)

32.0 (38)

=,=

Sargassum echinocarpum

10.0 (124)

10.0 (68)

10.0 (116)

=,=

32.0 (75)

32.0 (97)

10.0 (30)

=,↓

Sargassum polyphyllum

10.0 (33)

21.0 (54)

>54.0 (35)

>54.0 (36)

32.0 (42)

No Set 10.0 (36)

No Set 10.0 (98)

No Set >54.0 (36)

↑ ,↑ No Set

43.0 (60)

Sphacelaria tribuloides Turbinaria ornata

No Set 21.0 (33)

No Set 32.0 (30)

No Set 32.0 (44)

↑ ,↓ No Set

Rhodophyta Acanthophora spicifera

32.0 (49)

21.0 (60)

21.0 (35)

Asparagopsis taxiformis Galaxaura rugosa

No Set 10.0 (39)

No Set 10.0 (124)

No Set >54.0 (40)

Gracilaria coronopifolia

32.0 (98)

21.0 (57)

>54.0 (102)

Hypnea musciformis

10.0 (50)

32.0 (52)

54.0 (87)

Jania capillacea Kappaphycus alvarezii Laurencia cartilaginea

No Set No Set 21.0 (45)



Liagora tetrasporifera Melanamansia glomerata

No Set 21.0 (35)

No Set 21.0 (69)

No Set 10.0 (92)

Cyanobacteria Lyngbya majuscula Angiosperma Halophila hawaiiana

No Set

No Set

No Set

10.0 (31)

21.0 (59)

21.0 (64)

Control polystyrene

54 (30)

>54 (30)

>54 (30)

↑ ,↑ No Set

=,↑

21.0 (31)

>54.0 (46)

>54.0 (30)

=,↑

No Set >54.0 (91)

No Set >54.0 (62)

No Set >54.0 (53)

↓ ,↑

32.0 (54)

>54.0 (110)

32.0 (48)

↑ ,↑ No Set No Set

21.0 (45)

>54.0 (61)

>54.0 (61)

No Set No Set >54.0 (32)

No Set No Set >54.0 (32)


No Set >54.0 (64)

No Set >54.0 (46)

No Set 43.0 (46)

↓ ,= No Set

=,↑ No Set =,↓ No Set

=,=

No Set

↑ ,= ↑ ,= No Set =,= ↑ ,↓ ↑ ,= No Set No Set =,↓ No Set =,↓

No Set

No Set

No Set

↑ ,=

32.0 (38)

32.0 (42)

>54.0 (94)

No Set =,↑

↑ ,=

>54 (30)

>54 (30)

>54 (30)

=,=

which the plant tissues broke (Cr = 5.50: Colpomenia sinuosa, Acanthophora spicifera; Table 3). On 21 other plant species, H. elegans withstood less bending than the plant tissues (Fig. 3, Table 3). If individuals of H. elegans were removed when the diameter of the rod was greater than the length of their 7-d tubes (mean tube length ± S.E.: 3.6 ± 0.1 mm, N = 30), then individuals were always removed intact. On all remaining plant species, 7-d H. elegans were removed when bent around the 3.5 mm diameter rod (Cr = 5.50) either intact (six species), or in pieces (13 species) (Fig. 3). Thus, survival of H. elegans


823

Figure 2. Flow rates (cm s−1) required to remove 90% of 2-d, 7-d and 30-d Hydroides elegans from representative species of marine macrophytes. Examples where animal adhesion: (A) was consistent over time, (B) increased over time, (C) decreased over time, (D) increased then decreased over time and (E) a decrease was followed by an increase.

in the field is predicted to be limited to plants with limited flexibility (three species), or no flexibility (3 species; Fig. 3, Table 3). For 29 of 32 plant species, it was not possible to remove 90% of the 7-d B. neritina by bending plant tissues (Table 3). The attachment area of this arborescent bryozoan (0.40


824

Table 3. Flexibility of macrophytes and attached 7-d recruits of Hydroides elegans and Bugula neritina. Algal Flexibility = radius of curvature (Cr) required to break tissues of 90% of the tested plants (N ≥ 30). If Cr = 0, then it was not possible to break any plant tissues; if Cr = ∞ , then the plant tissues could not be bent without breaking. 90% Recruits Removed = Cr at which 90% of the animals were removed from the plant surface, either intact or in pieces. N.R. = not removed, with less than 90% of the recruits removed at the Cr at which the plant tissues broke; No Set = larval survival was significantly reduced or no larvae settled on the plant species, so flexibility trials were not run. Recruitment Potential = No, if ≥ 90% of the animals were removed at a Cr greater than the Cr at which the plant tissues broke, or Yes, if more than 10% of the animals remained attached to the plant at a Cr at which 90% of the plant tissues broke. Hydroides elegans Algal flexibility Chlorophyta Avrainvillea amadelpha Bornetella sphaerica Bryopsis pennata Caulerpa racemosa Caulerpa taxifolia Caulerpa verticillata Cladophora dotyana Codium arabicum Codium edule Codium reediae Dictyosphaeria cavernosa Enteromorpha flexuosa Halimeda discoidea Microdictyon setchellianum Neomeris annulata Ulva fasciata Ulva reticulata Valonia aegagropila Ventricaria ventricosa Phaeophyta Colpomenia sinuosa Dictyota acutiloba Dictyota crenulata Lobophora variegata Padina australis Sargassum echinocarpum Sargassum polyphyllum Sphacelaria tribuloides Turbinaria ornata Rhodophyta Acanthophora spicifera Asparagopsis taxiformis Galaxaura rugosa Gracilaria coronopifolia Hypnea musciformis Jania capillacea Kappaphycus alvarezii Laurencia cartilaginea Liagora tetrasporifera Melanamansia glomerata Cyanobacteria Lyngbya majuscula Angiosperma Halophila hawaiiana

0 ∞ 0 3.14 3.14 0 3.14 5.50 0 0 ∞ 0 7.85 7.85 5.50 0 0 12.57

90% recruits removed (N) 5.50 (30) N.R. (30) No set 5.50 (35) 5.50 (31) 5.50 (35) No Set 7.85 (38) 5.50 (68) 5.50 (30) N.R. (30)

Recruitment potential

pieces

No Yes

N.R. (38) N.R. (30)

Yes Yes

No No No No No No No No Yes

N.R. (50) N.R. (113) No Set N.R. (31) N.R. (44) N.R. (30) 1.57 (30) 1.57 (39) N.R. (30)

Yes Yes No Yes Yes Yes No No Yes

No No Yes No No No Yes Yes

N.R. (52) N.R. (30) N.R. (30) 5.50 (60) N.R. (59) N.R. (44) N.R. (30) N.R. (30)

No No No No No No No No Yes

No Set N.R. (38) N.R. (52) N.R. (44) N.R. (61) N.R. (30) N.R. (68) No Set N.R. (35)

No Yes Yes Yes Yes Yes Yes No Yes

No No No No No No No No No No

N.R. (33) No Set N.R. (30) N.R. (30) N.R. (64) No Set No Set N.R. (58) No Set N.R. (30)

Yes No Yes Yes Yes Yes No No No Yes

No

No Set

No

No

N.R. (70)

Yes

intact pieces pieces intact intact intact pieces

∞

5.50 (30) No Set N.R. (30) No Set 5.50 (30) 5.50 (30) N.R. (35) N.R. (30)

5.50 0 0 0 1.57 5.50 1.57 0 7.85

5.50 (30) 5.50 (32) No Set 5.50 (30) 5.50 (30) 10.21 (33) 5.50 (49) No Set N.R. (32)

pieces pieces

5.50 0.79 7.85 0.79 1.57 0.79 17.28 1.57 5.50 0

5.50 (32) No Set 10.21 (30) 5.50 (30) 5.50 (30) No Set No Set 5.50 (30) No Set 5.50 (30)

pieces

0

No Set

1.57

5.50 (69)

Bugula neritina

Nature of removal

pieces pieces

intact pieces intact pieces

intact pieces pieces

intact pieces

intact

90% recruits removed (N)

Nature of removal

intact intact

intact

Recruitment potential

Yes Yes Yes No Yes Yes Yes Yes

± 0.01 mm2; N = 30) was less than the thickness of many of the plant surfaces (L. J. Walters, per. obs.). Plant flexibility may, however, account for the absence of B. neritina on three species of green algae (Codium edule, C. reediae, Neomeris annulata) in the field (Table 3).


825

Figure 3. Relative flexibility of 7-d Hydroides elegans versus marine macrophytes when bent around rods of decreasing outer diameter (15.0–0.5 mm). If the radius of curvature (Cr) was 0, then the plant could not be broken using this protocol; if the radius of curvature = ∞, then the organism was totally inflexible. Representative examples where macrophytes (solid bars) were more flexible than 90% of tested H. elegans (diagonally striped bars) and the animals were removed (A) intact or (B) in fragments. Alternatively, some plants were less flexible than H. elegans and animals were not removed from these species (C).

826


DISCUSSION SUMMARY OF PRE- AND POST-SETTLEMENT FOULING DEFENSES.—If a submerged surface is free from fouling by sessile invertebrates in a habitat where space is limiting and a pool of competent larvae is present, then that surface is kept clean either by pre-settlement processes that deter larval attachment, or post-settlement processes that remove settled individuals. After determining that 40 species of Hawaiian marine plants were rarely fouled by sessile invertebrates, we evaluated the extent to which pre- and post-settlement processes were effective under controlled laboratory conditions; these results are summarized in Table 4. Pre-settlement mechanisms of anti-fouling were assigned if larval settlement on a plant was significantly reduced by toxicity or avoidance. Poor adhesion by the invertebrate was included as a post-settlement anti-fouling mechanism if 90% of the attached animals of any tested age were removed at a flow rate of 10 cm s−1. Although overall flow rates at all collection locations exceeded this value on a daily basis (Fig. 1; L. J. Walters, per. obs.), we have adopted this conservative estimate of the importance of this anti-fouling mechanism, as it is unlikely all algae would be subjected to the maximum recorded levels of water motion. Plant flexibility was included as a physical anti-fouling mechanism if 90% of the animals were removed from a plant species without the plant itself also breaking. With these considerations, one or more alternatives could explain the lack of H. elegans on all 40 plant species (Table 4). Pre-settlement processes could account for the lack of fouling on 23 plant species in our laboratory trials; post-settlement removal could explain the absence of H. elegans on the remaining 17 species. This differs significantly from man-made, submerged substrata in Hawaiian waters, which frequently have 100% cover of H. elegans (Edmundson, 1944; Walters et al., 1997). The lack of fouling by B. neritina can be explained by one of our tested alternatives for only 18 plant species (Table 4). For B. neritina, other alternatives, such as larval supply, abrasion or predation, may limit recruitment in Hawaiian waters. In Beaufort, North Carolina, Walters (1992a) found that fish consumed all juvenile and adult B. neritina not in spatial refuges. Fish were also observed to consume all accessible B. neritina in Hawaii where adults were collected (L. J. Walters, per. obs.). When results of survival and settlement-location preference trials for both sessile invertebrates were compared, some consistent patterns emerged (Table 1). Asparagopsis taxiformis and Lyngbya majuscula were toxic to both H. elegans and B. neritina. Caulerpa taxifolia, U. reticulata and Acanthophora spicifera were non-toxic but avoided by settling larvae of both species. The lack of toxicity with C. taxifolia was especially interesting in light of negative responses of many diverse marine organisms to the secondary metabolites associated with this species in the Mediterranean (e.g., Ferrer et al., 1997; Pedrotti and Lemee, 1999; Ricci et al., 1999; Uchimura et al. 1999). A positive response was found with Laurencia cartilaginea; both H. elegans and B. neritina preferentially settled on this red alga. The same anti-fouling mechanisms were successful in many cases when plants of the same genera were considered (Table 1). For example, with H. elegans, larval survival was high for the three tested species of Caulerpa, three species of Codium, two species of Ulva and two species of Sargassum. However, survival was 100% for Dictyota acutiloba, while only 57% for D. crenulata. With H. elegans, there were differences in settlementlocation preferences within two genera: Caulerpa and Codium. There were no differ-


827

Table 4. Summary of results from pre- and post-settlement mechanisms that deterred fouling in Hawaiian marine macrophytes. Pre-settlement anti-fouling mechanisms were included if exposure to a plant species either significantly reduced larval settlement or larvae significantly avoided settling on the plant surface. Limited attachment was included as a post-settlement anti-fouling mechanism if 90% of the animals of any age could be removed at a flow rate of 10 cm s−1. Plant flexibility was included if bending removed 90% of the attached animals without the plant tissues also breaking. Hydroides elegans Pre-Settlement Post-Settlement Toxic/Avoid Attachment Flexibility Chlorophyta Avrainvillea amadelpha Bornetella sphaerica Bryopsis pennata Caulerpa racemosa Caulerpa taxifolia Caulerpa verticillata Cladophora dotyana Codium arabicum Codium edule Codium reediae Dictyosphaeria cavernosa Enteromorpha flexuosa Halimeda discoidea Microdictyon setchellianum Neomeris annulata Ulva fasciata Ulva reticulata Valonia aegagropila Ventricaria ventricosa Phaeophyta Colpomenia sinuosa Dictyota acutiloba Dictyota crenulata Lobophora variegata Padina australis Sargassum echinocarpum Sargassum polyphyllum Sphacelaria tribuloides Turbinaria ornata Rhodophyta Acanthophora spicifera Asparagopsis taxiformis Galaxaura rugosa Gracilaria coronopifolia Hypnea musciformis Jania capillacea Kappaphycus alvarezii Laurencia cartilaginea Liagora tetrasporifera Melanamansia glomerata Cyanobacteria Lyngbya majuscula Angiosperma Halophila hawaiiana Control Polystyrene

Bugula neritina Pre-Settlement Toxic/Avoid

+

+

+ + +

+

Post-Settlement Attachment Flexibility

+ + + +

+

+ +

+ + + +

+ +

+ + +

+ + + +

+ +

+ + +

+ + +

+ + + +

+ +

+

+ +

+

+ + + +

+

+

+

+ + + + +

+ +

+ +

+ + + + +

+ + +

+ +

+ + +

+ +

+ + +

+

+ +

+

+

+ +

+

ences in survival within a plant genus in larval assays with B. neritina. Differences in settlement location of B. neritina did occur with species of Ulva, Caulerpa and Sargassum. FACTORS THAT INFLUENCE LARVAL SETTLEMENT ON HAWAIIAN PLANTS.—A chemical signal is often the primary cue in determining where a larva settles (Pawlik, 1992). To influence a larva, a chemical signal must be present on an exposed plant surface, or released into the water by the plant. Although laboratory assays may allow the accumulation of higher concentrations of chemicals than larvae would encounter in the field, especially if the creation of plant fragments induced production of secondary metabolites, chemicals likely explain the lack of settlement by H. elegans or B. neritina on some plants. For plant

828


species in which larval mortality was consistently high, it is likely that chemistry was involved. Additionally, differences in toxicity for H. elegans versus B. neritina also argue for chemical defenses for these algae. For example, survival of H. elegans was significantly reduced with Cladophora dotyana, Halimeda discoidea, Neomeris annulata, Dictyota crenulata, Kappaphycus alvarezii and Liagora tetrasporifera; survival of larvae of B. neritina was greater than 99% with these same macrophytes (Table 1). Alternatively, contact with the brown alga Sphacelaria tribuloides decreased survival of B. neritina, but not H. elegans. For the statistical analyses on survival and settlement-location preferences, data from plants collected from three locations on different dates were merged to maximize the chances of finding differences between trials and minimize the chances of identifying plant species with chemicals that consistently influenced larval survival or behaviors. Because collection and assay protocols were the same for all trials, any chemical released as the result of handling should be consistent for a species on any given collection date. Differences among trials within a plant species potentially resulted from several factors: (1) differences in the history of the plant either in terms of past damage (grazing pressure or frequency of physical disturbance events), (2) seasonality and phase in life-history for algae with isomorphic alternation of generations, or (3) differences in the physiological state for plants within a population (e.g., Van Alstyne, 1988, Arnold and Targett 2000). For species in which fragments were used in the assays, variability may also be the result of the fragment tested, as chemicals are not always uniformly distributed. Increased levels of defensive compounds may be found in reproductive tissues, regions immediately surrounding wounded tissue, actively growing regions of the plant, more exposed regions of plants, and outer meristem walls (e.g., Steinberg, 1984; Hay et al., 1988; Paul and Van Alstyne, 1988a,b; Carlson et al. 1989; Tugwell and Branch, 1989; Meyer and Paul, 1992). Fragments with scars or reproductive structures were not used in our assays, and similar amounts of new plant biomass were included in each dish. If exposure to a plant was not lethal to larvae in our settlement assays, but individuals avoided settling on these plants, then it was not possible with our experimental design to distinguish between chemical and physical anti-fouling mechanisms. The larvae may respond to non-toxic or sub-lethal deterrent compounds, or other features associated with the plant surfaces, including biofilms, wettability and topographic complexity (e.g., Sieburth and Conover, 1965; Davis et al., 1991; Walters et al., 1996). Schmitt et al. (1995) found that when larvae of B. neritina were placed in bowls with Dictyota menstrualis and Gracilaria tikvahiae, they contacted both species equally, but settled almost exclusively on G. tikvahiae. Thus, any short-term exposure to the surface of D. menstrualis was not toxic to these larvae. Longer exposure to high levels of the deterrent compounds associated with D. menstrualis did cause significant mortality and reduced the growth rates of survivors (Schmitt et al., 1995). Walters et al. (1996) found that water conditioned with six species of Hawaiian macroalgae (H. discoidea, U. reticulata, S. echinocarpum, S. polyphyllum, S. tribuloides, and L. cartilaginea) was not toxic, but significantly reduced the settlement frequency of larvae of H. elegans in replicate trials. In the present study, one of these algal species (H. discoidea) killed significant numbers of larvae of H. elegans when these larvae were exposed continuously to plant biomass. Settlement location was not random for the remaining plant species (Table 1). In conditioned-water trials with larvae of B. neritina, eight plant species significantly reduced settlement (Walters et al., 1996). Of these, long-term exposure to L. majuscula and S. tribuloides was toxic, while


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continuous exposure to U. fasciata and H. discoidea inhibited settlement on the algal surfaces. Unprotected surfaces in marine habitats are rapidly colonized by microbial communities, and these biofilms influence the settlement of larvae of many sessile invertebrates (e.g., ZoBell and Allen, 1935; Mitchell and Kirchman, 1984; Holmström and Kjelleburg, 1994; Hadfield, 1998). Hadfield et al. (1994) and Unabia and Hadfield (1999) have described the importance of bacterial films for settlement of H. elegans, and Maki et al. (1989) showed that settlement of B. neritina decreased as bacterial densities increased. Bacteria were present on the surfaces of all plant species tested (T. M. Michael, unpubl. data), precluding the absence of biological films as the reason why larvae avoided some surfaces. Likewise, none of the plant-surface wettabilities fell outside the acceptable range for settlement for these two invertebrates (R. E. Baier and A. E. Meyer, unpubl. data). Schmitt et al. (1995) also found that surface wettability was not responsible for the lack of settlement of B. neritina on D. menstrualis. Additionally, larvae of H. elegans do not actively seek out specific topographic features in still water (Walters et al., 1997). Settlement of B. neritina was, however, influenced by small-scale topographic complexity, with larvae settling in high numbers in crevice locations in both still and moving water (e.g., Walters, 1992a,b; Walters and Wethey, 1996; Walters et al., 1999). Thus, flat surfaces may be avoided by B. neritina, potentially explaining the lack of individuals on plants with limited topographic complexity, such as U. reticulata. Although there are numerous examples of marine plants inducing settlement of benthic invertebrates, positive chemical cues for settlement were rarely found in the present study with larvae of H. elegans (Crisp, 1976, 1984; Pawlik, 1992; Trowbridge and Todd 2001; Table 1). Walters et al. (1996) found that three algae, P. australis, U. fasciata and H. musciformis, released compounds that significantly stimulated settlement by H. elegans. In this study, we did not specifically examine the timing of settlement, but rather settlement location. Greater settlement on plant surfaces was not found on these three algal species (Table 1). In contrast, conditioned waters never increased the frequency of settlement over that in control dishes with larvae of B. neritina (Walters et al., 1996), although larvae of this species did settle more frequently than expected by chance on six plant species. Preferential settlement on plant surfaces included representatives of the green algae, brown algae, red algae, and the marine angiosperms. On all of these plants, larvae began to settle immediately after being added to the assay dishes, and many were partially metamorphosed by the time the final larva was added (≤ 2 min). In control dishes and in dishes with all other plant species, no settlement was observed during this initial period. ADHESION TO PLANT SURFACES.—If both H. elegans and B. neritina were easily removed from a plant, then the plant surface was non-adhesive. Alternatively, significant differences in attachment strength may be associated with morphological differences between the two invertebrates. Very little water motion or bending was required to remove both H. elegans and B. neritina from the three species of Codium and S. echinocarpum (Fig. 2, Table 2). Poor adhesion to Codium spp. was likely because animals attached to fine hairs, or utricles rather than to a continuous, smooth surface. Likewise, S. echinocarpum is covered with small, sub-surface crypts with many fine hairs; this arrangement may also have prevented secure attachment of epiphytes. The flow velocity necessary to remove H. elegans and B. neritina was very different for the remaining algae (Table 2). One explanation for this is that the area of attachment of H. elegans increased over time, while

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that of B. neritina remained constant. Denny (1988) found that small encrusting organisms were less tenacious than larger individuals of the same species. A second possibility involves differences in the adhesives released by the two invertebrates. The mucopolysaccarides that secure B. neritina to substrata are released only at metamorphosis, while serpulids secrete multiple adhesives throughout their lives (Crisp, 1984; CarpizoItuarte and Hadfield, 1998). The adhesives produced by B. neritina were very strong when individuals were tested two days after settlement, and all individuals remained attached to the control surface at the highest flow rate on all test dates. The flow rate necessary to remove 90% of the H. elegans on polystyrene increased from day 2 to day 7, then remained constant from day 7– day 30 (Table 2). The same pattern was found for H. elegans on many plants. The number of plants from which 90% of H. elegans could be removed at 10 cm s−1 declined from 17 species at day 2 to 8 species at day 7 and to 7 species at day 30. In some cases, we observed a decrease in the flow rate required to remove settlers of H. elegans and B. neritina as they aged. This was unexpected, especially for recruits of H. elegans, as they increased their attachment areas over time. Two alternatives may explain this decrease in flow rate. Sub-lethal chemical toxicity may have reduced viability of attached individuals over time. Schmitt et al. (1995) found that pachydictyl A from Dictyota menstrualis did not reduce larval survivorship in B. neritina, but did reduce mean growth rate of juveniles. Alternatively, if individuals settled in a spatial refuge, such as the crevices between cells of D. cavernosa, then these individuals initially experienced a reduced impact of water from the water-jet. With growth, however, more of the individuals projected beyond the refuge, making removal of H. elegans and B. neritina easier. It could be argued that all adhesion results are an underestimate of the invertebrates’ ability to adhere to the macrophyte surfaces because the animals were grown on macrophytes in a still water environment and attachment strength likely increases as flow rates increase. We agree with this and suggest that this makes our results even more conservative and impressive. With the growth conditions used for H. elegans, at some age individuals could not be removed from eight species of macroalgae or polystyrene. Likewise, B. neritina could not be removed from 22 macrophytes or polystyrene with the maximum tested flow rate of 54 cm s−1. If flow regime has a direct impact on the strength or surface area of basal attachment, then our results for adhesion studies may be underestimates of the abilities of both invertebrates to remain attached to substrata in field settings. Eyster and Pechenik (1987) found significant increases in strength of byssal attachment by the mussel Mytilus edulis with increased water agitation. However, there is no evidence that arborescent bryozoans or sessile tube-building polychaetes respond to increased flow with more or stronger adhesive or larger attachment areas. PLANT FLEXIBILITY.—Bugula neritina and H. elegans performed very differently in the flexibility trials. If individuals of H. elegans were not removed from the plant by bending the plant around rods of diameters greater than the length of individuals of H. elegans, then they were always removed when the rod diameter and the length of H. elegans were approximately equal (Fig. 3, Table 3). However, in many of the flexibility trials, it was not possible to remove B. neritina. Additionally, after removal from a plant, the probability of animal survival differed for H. elegans and B. neritina. Although individuals of B. neritina were always removed intact, it is unlikely that they would be able to attach to a new substratum as their only adhesive is released during metamorphosis (Crisp, 1984).


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However, individuals of H. elegans may survive and immediately begin attaching to a new surface when tubes are dislodged from a surface because many polychaetes actively resist removal from their tubes with hooked setae and adhesives are secreted throughout their lives (Wisely, 1958; Woodin and Merz, 1987; Carpizo-Ituarte and Hadfield, 1998). EVOLUTION AND DIVERSITY OF ANTI-FOULING MECHANISMS.—It is not known if chemical or physical anti-fouling protection evolved specifically to keep plants free from fouling, or if the apparent presence of these properties is coincidental. Many algal chemicals or secondary products are thought to have evolved to inhibit microbial growth or reduce herbivory (Paul, 1992). See Appendix A for list of chemicals extracted from the 40 tested plant species. Many herbivores respond to taste rather than smell, and many compounds that are known to reduce grazing are contained in membrane-bound vesicles and are released when the plant is damaged, while compounds that deter larval settlement must be either on the exposed surfaces of the plant or released in to the surrounding water (Paul, 1992; Schmitt et al., 1995; Walters et al., 1996). Some compounds successfully reduced both herbivory and larval settlement. For example, compounds produced by Halimeda spp. and Lyngbya majuscula appear to provide multiple functions (e.g., Paul and Van Alstyne, 1988a,b; Wylie and Paul, 1988). However, the effectiveness of chemicals associated with Caulerpa spp. on herbivores and fouling organisms is much more variable (e.g., Targett et al., 1986; Paul et al., 1987; Table 1). IMPLICATIONS FOR BIOLOGICAL FOULING CONTROL ON MAN-MADE SURFACES.—Although infrequently found on Hawaiian macrophytes, both H. elegans and B. neritina are important members of fouling communities that rapidly cover submerged man-made surfaces, including ship hulls, buoys, offshore platforms, and seawater intake lines of power plants; therefore, millions of dollars are spent each year to control fouling (Allen, 1953; Alberte et al., 1992). To date, copper and tin-based compounds have been used to reduce the recruitment of sessile invertebrates. Unfortunately, these compounds are toxic to many non-target and target species alike. By understanding how submerged living organisms, such as tropical macroalgae and seagrasses deter biological fouling, scientists may gain insight into environmentally sound ways to control this problem. Our results show that many Hawaiian marine plants successfully prevent fouling by sessile invertebrates by combining chemicals that deter settling larvae and post-settlement mechanisms that remove attached individuals. ACKNOWLEDGEMENTS We thank C. Unabia, T. Michael, I. Abbott, M. Denny, E. Bell, R. Woollacott, D. Wethey, V. Paul and N. Phillips for help with experimental designs and observations on marine flora and fauna of Hawaii, R. Li, P. Sacks, R. Chock, S. Sakata, and M. Lammers for field and laboratory assistance, and an anonymous reviewer for greatly improving the manuscript. This research was supported by Office of Naval Research Contract No. N00014-90-J-1932 to C. M. Smith, M. G. Hadfield, and I. A. Abbott.

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LITERATURE CITED Alberte, R. S., S. Snyder, B. J. Zahuranec and M. Whetstone. 1992. Biofouling research needs for the United States Navy: program history and goals. Biofouling 6: 91–95. Allen, F. E. 1953. Distribution of marine invertebrates by ships. Aust. J. Mar. Freshw. Res. 4: 307– 316. Arnold, T. M. and N. M. Targett. 2000. Evidence for metabolic turnover of polyphenolics in tropical brown algae. J. Chem. Ecol. 26: 1393–1410. Bak, R. P. M. and J. L. A. Borsboom. 1984. Allelopathic interaction between a reef coelenterate and benthic algae. Oecologia 63: 194–198. Bakus, G. J., N. M. Targett and B. Schulte. 1986. Chemical ecology of marine organisms: an overview. J. Chem. Ecol. 12: 951–987. Bell, E. C. and M. W. Denny. 1994. Quantifying “wave exposure”: a simple device for recording maximum velocity and results of its use at several field sites. J. Exp. Mar. Biol. Ecol. 181: 9–29. Bernstein, B. B. and N. Jung. 1979. Selective pressures and coevolution in a kelp canopy community in Southern California. Ecol. Monogr. 49: 335–355. Buss, L. W. 1990. Competition within and between encrusting clonal invertebrates. Trends Ecol. Evol. 5: 352–356. Cancino, J. M., J. Muñoz, M. Muñoz and M. C. Orellana. 1987. Effects of the bryozoan Membranipora tuberculata (Bosc.) on the photosynthesis and growth of Gelidium rex Santelices et Abbott. J. Exp. Mar. Biol. Ecol. 113: 105–112. Carlson, D. J., J. Lubchenco, M. A. Sparrow and C. D. Trowbridge. 1989. Fine-scale variability of lanosol and its disulfate ester in the temperate red alga Neorhodomela larix. J. Chem. Ecol. 15: 1321–1334. Carpizo-Ituarte, E. and M. G. Hadfield. 1998. Stimulation of metamorphosis in the polychaete Hydroides elegans Haswell (Serpulidae). Biol. Bull. 194: 14–24. Chadwick, N. E. 1988. Competition and locomotion in a free-living fungiid coral. J. Exp. Mar. Biol. Ecol. 123: 189–200. Crisp, D. J. 1976. Settlement responses in marine organisms. Pages 83–124 in R. C. Newell, ed. Adaptation to environments: essays on the physiology of marine animals. Butterworths, London. ________. 1984. Overview of research on marine invertebrate larvae, 1940–1980. Pages 103–126 in J. D. Costlow and R. C. Tipper, eds. Marine biodeterioration: an interdisciplinary study. Naval Institute Press, Annapolis. ________ and J. S. Ryland. 1960. Influence of filming and of surface texture on the settlement of marine organisms. Nature 185: 119. ________ and G. B. Williams. 1960. Effect of extracts from fucoids in promoting settlement of epiphytic Polyzoa. Nature 188: 1206–1207. Davis, A. R., A. J. Butler and I. VanAltena. 1991. Settlement behavior of ascidian larvae: preliminary evidence of inhibition by sponge allelochemicals. Mar. Ecol. Prog. Ser. 72: 117–123. __________, N. M. Targett, O. J. McConnell and C. M. Young. 1989. Epibiosis of marine algae and benthic invertebrates: natural products chemistry and other mechanisms inhibiting settlement and overgrowth. Pages 85–114 in P. J. Scheuer, ed. Bioorganic marine chemistry, vol. 3. SpringerVerlag, Berlin. Denny, M. W. 1988. Biology and mechanics of the wave-swept environment. Princeton University Press, Princeton. 329 p. de Nys, R., J. C. Coll and I. R. Price. 1991. Chemically mediated interactions between the red alga Plocamium hamatum (Rhodophyta) and the octocoral Sinularia cruciata (Alcyonacea). Mar. Biol. 108: 315–320. Edmundson, C. H. 1944. Incidence of fouling in Pearl Harbor. Occasional Papers of the Bernice P. Bishop Museum 18: 1–34.


833

Eldredge, L. G. and R. C. DeFelice. 2000. Checklist of the marine invertebrates of the Hawaiian Islands. http://www2.bishopmuseum.org/list_home.htm. Eyster, L. S. and J. A. Pechenik. 1987. Attachment of Mytilus edulis L. larvae on algal and byssal filaments is enhanced by water agitation. J. Exp. Mar. Biol. Ecol. 114: 99–110. Ferrer, E, A. Gomez-Garreta and M. A. Ribera. 1997. Effect of Caulerpa taxifolia on the productivity of two Mediterranean macrophytes. Mar. Ecol. Prog. Ser. 149: 279–287. Filion-Myklebust, C. and T. A. Norton. 1981. Epidermis shedding in the brown seaweed Ascophyllum nodosum (L.) LeJolis, and its ecological significance. Mar. Biol. Let. 2: 45–51. Gerhart, D. J., D. Rittschof and S. W. Mayo. 1988. Chemical ecology and the search for marine antifoulants. Studies in a predator-prey symbiosis. J. Chem. Ecol. 14: 1905–1917. Hadfield, M. G. 1998. The DP Wilson lecture: Research on settlement and metamorphosis of marine invertebrate larvae: past, present and future. Biofouling 12: 9–29. ____________, C. C. Unabia, C. M. Smith and T. M. Michael. 1994. Settlement preferences of the ubiquitous fouler Hydroides elegans. Pages 67–74 in M. F. Thompson, R. Nagabhushanam, R. Sarojini and M. Fingerman, eds. Recent developments in biofouling control. Oxford and IBH Publishing Company, New Delhi. Harvell, C. D. 1984. Predator-induced defense in a marine bryozoan. Science 224: 1357–1359. ___________, W. Fenical and C. H. Greene. 1988. Chemical and structural defenses of Caribbean gorgonians (Pseudopterogorgia spp.). I. Development of an in situ feeding assay. Mar. Ecol. Prog. Ser. 49: 287–294. Hay, M. E., Q. E. Kappel and W. Fenical. 1994. Synergisms in plant defenses against herbivores: interactions of chemistry, calcification, and plant quality. Ecology 75: 1714–1726. _________, V. J. Paul, S. M. Lewis, K. Gustafson, J. Tucker and R. Trindell. 1988. Can tropical seaweeds reduce herbivory by growing at night?: diel patterns of growth, nitrogen content, herbivory, and chemical versus morphological defenses. Oecologia 75: 233–245. Holmström, C. and S. Kjelleberg. 1994. The effect of external biological factors on settlement of marine invertebrate and new antifouling technology. Biofouling 8: 147–160. Johnson, C. R. and K. H. Mann. 1986. The crustose coralline alga, Phymatolithon Foslie, inhibits the overgrowth of seaweeds without relying on herbivores. J. Exp. Mar. Biol. Ecol. 96: 127– 146. Keats, D. W. and G. Maneveldt. 1994. Leptophytum foveatum Chamberlain & Keats (Rhodophyta, Corallinales) retaliates against competitive overgrowth by other encrusting algae. J. Exp. Mar. Biol. Ecol. 175: 243–251. Keough, M. J. 1986. The distribution of the bryozoan Bugula neritina on seagrass blades: settlement, growth and mortality. Ecology 67: 846–857. Lang, J. C. 1973. Interspecific aggression by scleractinian corals. II. Why the race is not always to the swift. Bull. Mar. Sci. 23: 260–279. Lindquist, N., M. E. Hay and W. Fenical. 1992. Defenses of ascidians and their conspicuous larvae: adult versus larval chemical defenses. Ecol. Monogr. 62: 547–568. Littler, M. M. and D. S. Littler. 1980. The evolution of thallus form and survival strategies in benthic marine macroalgae: field and laboratory tests of a functional form model. Am. Nat. 116: 25–44. __________, P. R. Taylor and D. S. Littler. 1983. Algal resistance to herbivory on a Caribbean barrier reef. Coral Reefs 2: 111–118. Lynch, W. F. 1947. The behavior and metamorphosis of the larvae of Bugula neritina (Linnaeus): experimental modification of the length of the free-swimming period and the responses to light and gravity. Biol. Bull. 92: 115–150. Magruder, W. H. and J. W. Hunt. 1979. Seaweeds of Hawaii. Oriental Publishing Company, Honolulu. 116 p. Maki, J. S., D. Rittschof, A. R. Schmidt, A. G. Snyder and R. Mitchell. 1989. Factors controlling attachment of bryozoan larvae: a comparison of bacterial films and unfilmed surfaces. Biol. Bull. 177: 295–302.

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Meyer, K. D. and V. J. Paul. 1992. Intraplant variation in secondary metabolite concentration in three species of Caulerpa (Chlorophyta: Caulerpales) and its effect on herbivorous fish. Mar. Ecol. Prog. Ser. 82: 249–257. Michael, T. M. and C. M. Smith. 1995. Lectins probe molecular films in biofouling: characterization of early films on non-living and living surfaces. Mar. Ecol. Prog. Ser. 119: 229–236. Mihm, J. W., W. C. Banta and G. I. Loeb. 1981. Effects of adsorbed organic and primary fouling films on bryozoan settlement. J. Exp. Mar. Biol. Ecol. 54: 167–179. Mitchell, R. and D. Kirchman. 1984. The microbial ecology of marine surfaces. Pages 49–56 in J. D. Costlow, and R. C. Tipper, eds. Marine biodeterioration: an interdisciplinary study. Naval Institute Press, Annapolis. Muñoz, J., J. M. Cancino and M. X. Molina. 1991. Effect of encrusting bryozoans on the physiology of their algal substratum. J. Mar. Biol. Assoc. U.K. 71: 877–882. Paul, V. J. 1992. Seaweed chemical defenses on coral reefs. Pages 24–50 in V. J. Paul, ed. Ecological roles of marine natural products. Comstock Publishing Association, Ithaca. ________, M. M. Littler, D. S. Littler and W. Fenical. 1987. Evidence for chemical defense in tropical green alga Caulerpa ashmeadii (Caulerpaceae: Chlorophyta): isolation of new bioactive sesquiterpenoids. J. Chem. Ecol. 13: 1171–1185. ________ and K. L. Van Alstyne. 1988a. Activation of chemical defenses in the tropical green algae Halimeda spp. J. Exp. Mar. Biol. Ecol. 160: 191–203. ________ and ________________. 1988b. Chemical defense and chemical variation in some tropical Pacific species of Halimeda (Halimedaceae; Chlorophyta). Coral Reefs 6: 263–269. Pawlik, J. R. 1992. Chemical ecology of the settlement of benthic marine invertebrates. Oceanogr. Mar. Biol. Annu. Rev. 30: 273–335. Pedrotti, M. L. and R. Lemee. 1999. Effect of microalgae treated with natural toxins on the nutrition and development of filter-feeding sea-urchin larvae. Mar. Environ. Res. 48:177–192. Pennings, S. C. and V. J. Paul. 1992. Effect of plant toughness, calcification, and chemistry on herbivory by Dolabella auricularia. Ecology 73: 1606–1619. Porter, J. W. and N. M. Targett. 1988. Allelochemical interactions between sponges and corals. Biol. Bull. 175: 230–239. Ricci, N., Capovani, C. H., and F. Dini. 1999. Behavioral modifications imposed to the ciliate protist Euplotes crassus by caulerpenyne: The major toxic terpenoid of the green seaweed, Caulerpa taxifolia. Eur. J. Protistol. 35: 290–303. Rittschof, D., I. R. Hooper and J. D. Costlow. 1988. Settlement inhibition of marine invertebrate larvae: comparison of sensitivities of bryozoan and barnacle larvae. Pages 599–608 in M. F. Thompson, R. Sarojini and R. Nagabhushanam, eds. Marine biodeterioration. Oxford and IBH Publishing Company, New Delhi. Russell, G. and V. J. Veltkamp. 1984. Epiphyte survival on skin-shedding macrophytes. Mar. Ecol. Prog. Ser. 18: 149–153. Sammarco, P. W., J. C. Coll and S. La Barre. 1985. Competitive strategies of soft corals (Coelenterata: Octocorallia): II. Variable defensive responses and susceptibility to scleractinian corals. J. Exp. Mar. Biol. Ecol. 91: 199–215. SAS Institute. 1988. SAS user’s guide: statistics, 6.03 Edition. SAS Institute, Incorporated, Cary. Schmitt, T. M., M. E. Hay and N. Lindquist. 1995. Constraints on chemically mediated coevolution: multiple functions for seaweed secondary metabolites. Ecology 76: 107–123. Seed, R. 1986. Ecological pattern in the epifaunal communities of coastal macroalgae. Pages 22– 35 in P. G. Moore, and R. Seed, eds. The ecology of rocky coasts. Columbia University Press, New York. ______ and R. J. O’Connor. 1981. Community organization in marine algal epifaunas. Annu. Rev. Ecol. Syst. 12: 49–74. Sieburth, J. M. and J. T. Conover. 1965. Sargassum tannin, an antibiotic which retards fouling. Nature 208: 52–53.


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Smith, C. M. and L. J. Walters. 1999. Vegetative fragmentation in three species of Caulerpa (Chlorophyta, Caulerpales): The importance of fragment origin, fragment length, and wound dimensions as predictors of success. P.S.Z.N. Mar. Ecol. 20: 307–319. Sokal, R. R. and F. J. Rohlf. 1995. Biometry, 3rd ed. Freeman, San Francisco. 887 p. Steinberg, P. D. 1984. Algal chemical defense against herbivores: allocation of phenolic compounds in the kelp Alaria marginata. Science 223: 405–406. Targett, N. M., T. E. Targett, N. H. Vrolijk and J. C. Ogden. 1986. Effect of macrophyte secondary metabolites on feeding preferences of the herbivorous parrotfish Sparisoma radians. Mar. Biol. 92: 141–148. Todd, J. S., R. C. Zimmerman, P. Crews and R. S. Alberte. 1993. The antifouling activity of natural and synthetic phenolic acid sulfate esters. Phytochemistry 34: 401–404. Trowbridge, C. D. 1993. Interactions between an ascoglossan sea slug and its green algal host: branch loss and the role of epiphytes. Mar. Ecol. Prog. Ser. 101: 263–272. ______________. and C. D. Todd 2001. Host-plant change in marine specialist herbivores: Ascoglossan sea slugs on introduced macroalgae. Ecol. Monogr. 71: 219–243. Tugwell, S. and G. M. Branch. 1989. Differential polyphenolic distribution among tissues in the kelps Laminaria pallida and Macrocystis angustifolia in relation to plant-defense theory. J. Exp. Mar. Biol. Ecol. 129: 219–230. Uchimura, M., R. Sandeaux and C. Larroque. 1999. The enzymatic detoxifying system of a native Mediterranean scorpion fish is affected by Caulerpa taxifolia in its environment. Environ. Sci. Technol. 33: 1671–1674. Unabia, C. R. C. and M. G. Hadfield. 1999. Role of bacteria in larval settlement and metamorphosis of the polychaete Hydroides elegans. Mar. Biol. 133: 55–64. Van Alstyne, K. L. 1988. Herbivore grazing increases polyphenolic defenses in the intertidal brown alga Fucus distichus. Ecology 69: 655–663. ______________ and V. J. Paul. 1992. Chemical and structural defenses in the sea fan Gorgonia ventalina: effects against generalist and specialist predators. Coral Reefs 11: 155–159. Wahl, M. 1989. Marine epibiosis. I. Fouling and anti-fouling: some basic aspects. Mar. Ecol. Prog. Ser. 58: 175–189. Walters, L. J. 1992a. Post-settlement success of the arborescent bryozoan Bugula neritina (L.): the importance of structural complexity. J. Exp. Mar. Biol. Ecol. 164: 55–71. __________. 1992b. Field settlement locations on subtidal marine hard substrata: is active larval exploration involved? Limnol. Oceanogr. 37: 1101–1107. ___________, M. G. Hadfield and C. M. Smith. 1996. Waterborne chemical compounds in tropical macroalgae: positive and negative cues for larval settlement. Mar. Biol. 126: 383–393. ___________, ____________ and K. A. del Carmen. 1997. The importance of larval choice and hydrodynamics in creating aggregations of Hydroides elegans (Polychaeta: Serpulidae). Invert. Biol. 116: 102–114. ____________, G. Miron and E. Bourget. 1999. Endoscopic observations of invertebrate larval substratum exploration and settlement. Mar. Ecol. Prog. Ser. 182: 95–108. ____________ and C. M. Smith. 1994. Rapid rhizoid production in Halimeda discoidea Decaisne (Chlorophyta, Caulerpales) fragments: a mechanism for survival after separation from adult thalli. J. Exp. Mar. Biol. Ecol. 175: 105–120. ____________ and D. S. Wethey. 1991. Settlement, refuges, and adult body form in colonial marine invertebrates: a field experiment. Biol. Bull. 180: 112–118. ____________ and ___________. 1996. Settlement and early post-settlement survival of sessile marine invertebrates on topographically complex surfaces: the importance of refuge dimensions and adult morphology. Mar. Ecol. Prog. Ser. 137: 161–171. Williams, G. B. 1964. The effect of extracts of Fucus serratus in promoting the settlement of larvae of Spirorbis borealis (Polychaeta). J. Mar. Biol. Assoc. U.K. 44: 397–414. Wisely, B. 1958. The development and settling of a serpulid worm, H. elegans norvegica Gunnerus (Polychaeta). Aust. J. Mar. Freshwater Res. 9: 351–361.

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Woodin, S. A. and R. A. Merz. 1987. Holding on by their hooks: anchors for worms. Evolution 41: 427–432. Woollacott, R. M. and W. J. North. 1971. Bryozoans of California and North Mexico kelp beds. Beih. Nova Hedwigia 32: 455–479. Wylie, C. R. and V. J. Paul. 1988. Feeding preferences of the surgeonfish Zebrasoma flavescens in relation to chemical defenses of tropical algae. Mar. Ecol. Prog. Ser. 45: 23–32. ZoBell, C. E. and E. C. Allen. 1935. The significance of marine bacteria in the fouling of submerged surfaces. J. Bact. 29: 239–251. DATE SUBMITTED: June 22, 2001.

DATE ACCEPTED: May 24, 2002.

ADDRESSES: (L.J.W., C.M.S.) University of Hawaii, Department of Botany, 3190 Maile Way, Honolulu, Hawaii 96822. (L.J.W., M.G.H.) University of Hawaii, Kewalo Marine Laboratory, 41 Ahui Street, Honolulu, Hawaii 96813. CORRESPONDING AUTHOR: (L.J.W.) Department of Biology, University of Central Florida, Orlando, Florida 32816. E-mail: .

None described None described Bryopsin1 Caulerpenyne1,2; Caulerpicin3,4; Caulerpin1,4,5; Caulerpin analogues6; Cinnamyl dihydrocinnamate5; Cinnamyl hydrocinnamate7; Cinnamyl-3-phenyl-1-propenyl ether5; trans-cinnamyl-1-phenyl-2-propenyl ether5; Dicinnamyl ether5; 1,5diphenyl-1,4-pentadiene5; Iodine8; 10-keto-2,7,11-trimethyldodecanoic acid6 Caulerpa taxifolia Caulerpenyne2; Clionasterol9; Cholesterol9; 24-ethylidene-cholesterol9; 24-methyl-cholesterol9; 24-methylene-cholesterol9 Caulerpa verticillata Caulerpenyne2 Cladophora dotyana None described Codium arabicum Clerosterol10; (24S)-24-ethyl-5-a-hydroperoxycholesta-6,25-dien-3-b-ol10; (24S)-24-ethyl-7-a-hydroperoxycholesta-5,25-dien3-b-ol10; (24S)-24-ethyl-7-oxocholesta-5,25-dien-3-b-ol10; (24S)-24-ethylcholesta-5,25-dien-3b,7a-diol10; (24S)-24oxocholesta-4,25-dien-6-b-ol10 Codium edule None described Codium reediae None described Dictyosphaeria cavernosa Ergosterol11 Enteromorpha flexuosa Iodine8; Palmitic acid12; Sulphonoglycolipid13 Halimeda discoidea Halimedatetracetate1; Halimedatrial1 Microdictyon setchellianum None described Neomeris annulata Brominated sesquiterpenes14; Neomeranol15 Ulva fasciata Erythro-octadecasphinga-4,8-dienine-N-palmitate16; Iodine8; Isofucosterol17; 2-N-palimitoyl-4,5-dihydro-1,3,4,5-tetrahydroxy sphingosine17; Sarringosterol17; b-sitosterol17 Ulva reticulata None described Valonia aegagropila None described Ventricaria ventricosa None described

Chlorophyta Avrainvillea amadelpha Bornetella sphaerica Bryopsis pennata Caulerpa racemosa

Appendix A Known secondary metabolites in Hawaiian macrophytes used in our study


837

Liagora tetrasporifera Melanamansia glomerata

Jania capillacea Kappaphycus alvarezii Laurencia cartilaginea

Galaxaura rugosa Gracilaria coronopifolia Hypnea musciformis

Asparagopsis taxiformis

Lobophora variegata Padina australis Sargassum echinocarpum Sargassum polyphyllum Sphacelaria tribuloides Turbinaria ornata Rhodophyta Acanthophora spicifera

Dictyota acutiloba Dictyota crenulata

Phaeophyta Colpomenia sinuosa

None described 24-methylene cholesterol49

Arachidic acid32; Behenic (C22) acid32; Carrageenan33; Cholest-4-ene-3α,6β-diol34; Cholest-4-ene-3-one34; Cholesterol32; Iodine8; Lauric acid34; Methyl palmitate32; O-pthalic acid bis-(2 ethylnonyl)-ester34; Palmitic acid32; Stearic acid32 Bromoform35; Dibromochloromethane36; Dibromomethane36; 1,2-dibromomethylene36; Dihaloacetic acid35; Dihaloacrylic acid35; Haloacetic acid35; Haloacrylic acid35; Iodine8; Perchloroethylene37; Tribromoethylene36; Trichloroethylene37; Trihaloacrylic acid35 None described Anhydrodebromoaplysiatoxin38; Aplysiatoxin39; Debromoaplysiatoxin39; Manauealide C38 Carrageenan40; Kappa-carrageenan41; 5b-cholest-1-ene-20-hydroxy-7,11-diene42; 5b-cholest-3-ene-7,11-dione (Hypneadione)43,44; Cholesterol45; Iodine8; Isothionic acid46 None described Kappa-carrageenan47 Aplysistatin (stated as metabolite in all species of Laurencia)23; Elatol (stated as metabolite in all species of Laurencia)23; Floridoside (stated as metabolite in all species of Laurencia)48; Palisadan (stated as metabolite in all species of Laurencia)23

Brassicasterol18; Campesterol18; Cholesterol18; Clionasterol18; Colpol19; 22-(E)dehydrocholesterol18; Desmosterol18; Fucosterol18; 24-methylene cholesterol18; Methyl benhate20; Methyl lignocerate20; Methyl myristate20; Methyl oleate20; Methyl palmitate20; Methyl tetradecatrienoate20; Phosphatidylcholine21; Poriferasterol18 Acutilol A22; Acutilol A acetate22; Acutilol B22; Pachydictyol A (stated as metabolite in all species of Dictyota)23 Acetoxycrenulide24; β-Crenulal25; 18,0-dihydro-4β-hydroxydictyodial A-18-acetate26; 4β-hydroxydictyodial A26; Pachydictyol A (stated as metabolite in all species of Dictyota)23 Orcinol27; Phloroglucinol27; Phlorotannins28; Salicylic Acid27; Usnic Acid27 Alginic acid29 None described None described None described Alginic acid29; Alginin30; 20-hydroxy-4,8,13,17-tetramethyl-4,8,12,16-eicosatetraenoic acid31; Mannitol30

Appendix A (continued) Known secondary metabolites in Hawaiian macrophytes used in our study

838 BULLETIN OF MARINE SCIENCE, VOL. 72, NO. 3, 2003

None described

Antillatoxin50,51; Aplysiatoxin52; Barbaramide A50; α-Butyrolactone53; Carmabin A54; Carmabin B54; Curacin A55,56,57; Curacin B55; Curacin C55; Curacin D55; Curacin E55; Curacin F55; Debromoaplysiatoxin52,58,59; 4,8-dimethyl-6-0-(2',4'-di-0-methylβDxylopyranosyl)-hydroxyquinoline60; 4,8-dimethyl-6-hydroxyquinoline60; N,7-dimethylindole-3-carboxaldehyde53; Dolastatin61; Granadadiene54; Granadamide54; Hermitamide A62; Hermitamide B62; 5-hydroxymethyl-5-benzyl-valerolactone63; Kalkipyrone54; Kalkitoxin64; Kulolide65; Kulokainalide65; Lyngbyacarbonate55; Lyngbyastatin61; Lyngbyatoxin66; Lyngbyatoxin A67,68,69; Lyngbyatoxin B70; Lyngbyatoxin C70; Majusculamide A71; Majusculamide B71; Majusculamide C72; Malyngamide A1,73,74; Malyngamide B1,73,74; Malyngamide I54; Malyngamide J54; Malyngamide K54; Malyngamide L54; Malyngamide Q75; Malyngamide R75; Malyngamide B acetate73; Malyngamide H51,76; Malyngolide74; (4E,7S)-(-)-7-methoxy-4-tetradecenoic acid77; 7-methoxytetradec-4-enoic acid76; 5-methyl-3-pyrolin-2-one protein78; Methyl tumonoate A79; Methyl tumonoate B79; Microcolin A78,80; Microcolin B80; Tumonoic acid A79, Tumonoic Acid B79; Tumonoic acid C79; Yanucamide A65; Yanucamide B65; Ypaoamide81

1 Paul, V. J., S. G. Nelson and H. R. Sanger. 1990. Mar. Ecol. Prog. Ser. 60: 23−34; 2Paul, V. J. and W. Fenical. 1986. Mar. Ecol. Prog. Ser. 34: 157−169; 3Nielsen, P. G., J. S. Carle and C. Christopherson. 1982. Phytochemistry 21: 1643−1645; 4Vest, S. E., C. J. Dawes and J. T. Romeo. 1983. Bot. Mar. 26: 313−316; 5Anjaneyulu, A. S. R., C. V. S. Prakash, K. V. S. Raju and U. V. Mallavadhani. 1992. J. Nat. Prod. 55: 496−499; 6Anjaneyulu, A. S. R., C. V. S. Prakash and U. V. Mallavadhani. 1991. Phytochemistry 22: 3041−3042; 7Subbaraju, G. V. and D. Rajasekhar. 1996. Indian J. Chem. 35: 615−616; 8Mairh, O. P., B. K. Ramavat, A. Tewari, R. M. Oza and H. V. Joshi. 1989. Phytochemistry 28: 3307−3310; 9Valls, R., J. Artaud, A. Archavlis, N. Vicente and L. Piovetti. 1994. Oceanol. Acta 17: 223−226; 10Sheu, J-H, C-C Liaw and C-Y Duh. 1995. J. Nat. Prod. 58: 1521−1526; 11Al Easa, H. S., J-M Kornprobst, and A. M. Rizk. 1995. Phytochemistry 39: 373−374; 12Parekh, R. G., H. H. Parekh, and P. S. Rao. 1984. Indian J. Mar. Sci. 13: 45−46; 13Siddhanta, A. K., B. K. Ramavat, V. D. Chauhan, B. Achari, P. K. Dutta and S. C. Pakrashi. 1991. Bot. Mar. 34: 365−367; 14Meyer, K. D. and V. J. Paul. 1995. Mar. Biol. 122: 537−545; 15Barnekow, D. E., J. H. Cardellina II, A. S. Zektzer and G. E. Martin. 1989. J. Am. Chem. Soc. 111: 3511−3517; 16Sharma, M., H. S. Garg and K. Chandra. 1996. Bot. Mar. 39: 213−215; 17Pandey, M., H. S. Garg, D. S. Bhakuni and M. M. Dhar. 1991. Pages 283−287 in M-F Thompson, R. Sarojini and R. Nagabhushanam, eds. Bioactive Compounds from Marine Organisms with Emphasis on the Indian Ocean; 18Aknin, M., K. Dogbevi, A. Samb, J-M Kornprobst, E. M. Gaydou, and J. Miralles. 1992. Comp. Biochem. Physiol. 102B: 841−843; 19 Green, D., Y. Kashman and A. Miroz. 1993. J. Nat. Prod. 56: 1201−1202; 20Shakih, W., M. Shameel, V. U. Ahmad and K. Usmanghani. 1991. Bot. Mar. 34: 77−79; 21Araki, S., W. Eichenberger, T. Sakurai and N. Sato. 1991. Plant Cell Physiol. 32: 623−628; 22Hardt, I. H., W. Fenical, G. Cronin and M. E. Hay. 1996. Phytochemistry 43: 71−73; 23Paul, V. J., C. R. Wylie and H. R. Sanger. 1988. Proceedings of the Sixth International Coral Reef Symposium 3: 73−78; 24Sun, H. H., F. J. McEnroe and W. Fenical. 1983. J. Org. Chem. 48: 1903−1906; 25Kirkup, M. P. and R. E. Moore. 1983. Phytochemistry 22: 2527−2529; 26Kirkup, M. P. and R. E. Moore. 1983. Phytochemistry 22: 2539−2541; 27Legaz, M. E., C. Vicente and L. Xavier Filho. 1985. Cryptogamie: Algol. 6: 265−272; 28Stern, J. L., A. E. Hagerman, P. D. Steinberg and P. K. Mason. 1996. J. Chem. Ecol. 22: 1877−1899; 29Laserna, E. C., R. L. Veroy, A. H. Luistro, N. E. Montano and G. J. B. Cajipe. 1982. Kalikasan 11: 51−56; 30Kallaperumal, N., S. Kalimuthu and J. R. Ramalingam. 1989. J. Mar. Biol. Assoc. India 31: 303−305; 31Sawai, Y., Y. Fujita, K. Sakata and E. Tamashiro. 1994. Fish. Sci. 60: 199−201; 32 Wahidulla, S., L. D. Souza and S. Y. Kamat. 1986. Bot. Mar. 29: 49−50; 33Parekh, R. G., Y. A. Doshi and V. D. Chauhan. 1989. Indian J. Mar. Sci. 18: 139−140; 34Wahidulla, S., L. D'Souza and M. Govenker. 1998. Phytochemistry 48: 1203−1206; 35Woolard, F. X., R. E. Moore and D. P. Roller. 1979. Phytochemistry 18: 617−620; 36Marshall, R. A., D. B. Harper, W. C. McRoberts and M. J. Dring. 1999. Limnol. Oceanogr. 44: 1348−1352; 37Abrahamson K. and A. Ekdahl. 1995. Limnol. Oceanogr. 40: 1321−1326; 38Nagai, H, Y. Kan, T. Fujita, B. Sakamoto and Y. Hokama. 1998. Biosci., Biotechnol. Biochem. 62: 1011−1013; 39Nagai, H., T. Yasumoto and Y. Hokama. 1996. Toxicon 34: 753−761; 40Castro Solano, J. M. , and R. Corella Vargas. 1992. Brenesia 37: 59−66; 41Parekh, R. G., Y. A. Doshi, A. V. Rao and V. D. Chauhan. 1988. Indian J. Mar. Sci. 17:242−243; 42Babu, J. M., G. K. Trivedi, and H. H. Mathur. 1990. Phytochemistry 29: 2029−2031; 43Babu, J. M., G. K. Trivedi and H. H. Mathur. 1989. Phytochemistry 28: 3237−3239; 44Moses, J. B., G. K. Trivedi and H. H. Mathur. 1991. Pages 195−200 in M-F Thompson, R. Sarojini and R. Nagabhushanam, eds. Bioactive compounds from marine organisms with emphasis on the Indian Ocean; 45Afaq-Hussein, S., M. Shameel and R. Khan. 1992. Bot. Mar. 35: 141−146; 46Holst, P. B., S. E. Nielsen, U. Anthoni, K. S. Bisht, C. Christopherson, S. Gupta, V. S. Parmar, P. H. Nielsen and D. B. Sahoo. 1994. J. Appl. Phycol. 6: 443−446; 47Santos, G. A. 1989. Aquat. Bot. 36: 55−67; 48Barrow, K. D., Karsten, U., King, R. J. and J. A. West. 1995. Phycologia 34: 279−283; 49Combaut, G., L. Codomier, J. Teste and M. Pedersen. 1981. Phytochemistry 20: 1748; 50Orjala, J., D. Nagle and W. H. Gerwick. 1995. J. Am. Chem. Soc. 117: 8281−8282; 51Orjala, J., D. Nagle and W. H. Gerwick. 1995. Toxicology: Toxin Rev. 14: 109; 52Moore, R. E. 1984. Am. Chem. Soc. Sympos. Ser. 262: 369−376; 53Todd, J. S. and W. H. Gerwick. 1995. J. Nat. Prod. 58: 586−589; 54Nagle, D. G. and V. J. Paul. 1999. J. Phycol. 35: 1412−1421; 55Gerwick, W. H., H-D Yoo, D. Nagle, J. Orjala, M. A. Roberts, P. Protean, E. Hamel and A. Blokhin. 1995. Toxin Rev. 14: 110; 56White, J. D., T-S Kim and M. Nambu. 1995. J. Am. Chem. Soc. 117: 5612−5613; 57White, J. D. , T-S Kim and M. Nambu. 1997. J. Am. Chem. Soc. 119: 103−111; 58Solomon, A. E. and R. B. Stoughton. 1978. Arch. Dermatology 114: 1333−1335; 59Beutler, J. A., A. B. Alvarado, D. E. Schaufelberger, P. Andrews and T. G. McCloud. 1990. J. Nat. Prod. 53: 867−874; 60Orjala, J. and W. H. Gerwick. 1997. Phytochemistry 45: 1087−1090; 61Harrigan, G. G., H. Luesch, W. Y. Yoshida, R. E. Moore, D. G. Nagle, J. Biggs, P. U. Park and V. J. Paul. 1999. J. Nat. Prod. 62: 464−467; 62Tan, L. T., T. Okino and W. H. Gerwick. 2000. J. Nat. Prod. 63: 952−955; 63Zeng, H. 1994. Chin. J. Mar. Drugs 13: 31−33; 64Berman, F. W., W. H. Gerwick and T. F. Murray. 1999. Toxicon 37: 1645−1648; 65Sitachitta, N., R. T. Williamson and W. H. Gerwick. 2000. J. Nat. Prod. 63: 197−200; 66Moore, R. E. 1977. Bioscience 27: 797−802; 67Robinson, C. P., D. R. Franz and M. E. Bondura. 1991. Toxicon 29: 1009−1017; 68Stafford, R. G., M. Menta, and B. W. Kemppainen. 1992. Food Chem. Toxicol. 30: 795−801; 69Cardellina II, J. H., F-J Marner and R. E. Moore. 1990. Science (Wash.) 38: 2847−2849; 70Aimi, N., H. Odaka, S-I Sakai, H. Fujiki, M. Suganuma, R. E. Moore and G. M. L. Patterson. 1990. J. Nat. Prod. 53: 1593−1596; 71Marner, F-J, R. E. Moore, K. Hirotsu and J. Clardy. 1977. J. Org. Chem. 42: 2815−2819; 72Moore, R. E. 1996. J. Indian Microbiol. 16: 134−143; 73Pennings, S. C., A. M. Weiss and V. J. Paul. 1996. Mar. Biol. 126: 735−743; 74Thacker, R. W., D. G. Nagle and V. J. Paul. 1997. Mar. Ecol. Prog. Ser. 147: 21−29; 75Milligan, K. E., B. Marquez, R. T. Williamson, M. Davies-Coleman and W. H. Gerwick. 2000. J. Nat. Prod. 63: 965−968. 76Orjala, J., D. Nagle and W. H. Gerwick. 1995. J. Nat. Prod. 58: 764−768; 77Mueller, C., G. Voss and H. Gerlach. 1995. Liebigs Annuals 1995: 673−676; 78Koehn, F. E., O. J. McConnell, R. E. Longley, S. H. Sennett and J. K. Reed. 1994. J. Med. Chem. 39: 3181−3186; 79Harrigan, G. G., H. Luesch, W. Y. Yoshida, R. E. Moore, D. G. Nagle, J. Biggs, P. U. Park and V. J. Paul. 1999. J. Nat. Prod. 62: 464−467; 80Koehn, F. E., R. E. Longley and J. K. Reed. 1992. J. Nat. Prod. 55: 613−619; 81Nagle, D. G. and V. J. Paul. 1998. J. Exp. Mar. Biol. Ecol. 225: 29−38.

Angiosperma Halophila hawaiiana

Cyanobacteria Lyngbya majuscula

Appendix A (continued) Known secondary metabolites in Hawaiian macrophytes used in our study


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