Habitat Selection of Coral Reef Invertebrates

HABITAT SELECTION OF CORAL REEF INVERTEBRATES

MEDIATED BY ENVIRONMENTAL STIMULI

Molly M. Ashur

A dissertation submitted to the Faculty of the University of Delaware in partial fulfillment of the requirements for the degree of Doctor of Philosophy in Marine Studies

Summer 2019

HABITAT SELECTION OF CORAL REEF INVERTEBRATES

MEDIATED BY ENVIRONMENTAL STIMULI

Molly M. Ashur

Approved: ______Mark Moline, Ph.D. Chair of the Department of Marine Science and Policy

Approved: ______Estella Atekwana, Ph.D. Dean of the College of Earth, Ocean and Environment

Approved: ______Douglas J. Doren, Ph.D. Interim Vice Provost for Graduate and Professional Education

I certify that I have read this dissertation and that in my opinion it meets the academic and professional standard required by the University as a dissertation for the degree of Doctor of Philosophy.

Signed: ______Danielle L. Dixson, Ph.D. Professor in charge of dissertation

Signed: ______Mark E. Warner, Ph.D. Member of dissertation committee

Signed: ______Jonathan H. Cohen, Ph.D. Member of dissertation committee

Signed: ______Valerie Paul, Ph.D. Member of dissertation committee

ACKNOWLEDGMENTS

There are so many people who made this dissertation possible. First of all, thank you to my advisor, Dr. Danielle Dixson, for her guidance, support, encouragement, and pep talks throughout this journey and for all of the incredible opportunities she’s given me. A huge thanks to my committee members, Dr. Mark Warner and Dr. Jon Cohen, for critiques and guidance throughout my many projects. Thank you also to my external committee member, Dr. Valerie Paul, not only for research support and mentorship, but also for cheering me on in the field. I would like to thank the wonderful staff at the University of Delaware for all their administrative support, as well as the researchers and staff at the Smithsonian Institution in Belize for their encouragement and camaraderie. Vinaka vakalevu to my Fijian family at Coralview, Matava, and Barefoot Manta for making Fiji feel like home during research trips. I would also like to acknowledge all of the funding sources that made this research possible: NSF, NIH, Gordon and Betty Moore Foundation, Association of Marine Laboratories of the Caribbean, Georgia Institute of Technology, and University of Delaware. Words can’t express how grateful I am to have been a part of such an incredible lab. A special thank you to Rohan Brooker, who was there at the very start at Georgia Tech and has been a constant mentor throughout. Lane Johnston, the Ann to my Leslie, thank you for your friendship, spirit, and love through the ups and downs. A massive shout out to my other field mates: Paul, JJ, Alex, and Zara. Thanks for dealing with all the crazy and keeping me sane in the field. To the rest of the lab,

Emily, Taylor, Matt, Will, and the many interns who played a huge role in helping me accomplish this research, thank you so much for your encouragement, support, and laughter throughout this whole process. I would also like to thank my family, near and far. Thank you to my wonderful in-laws who welcomed me with open arms and continually encouraged me. Thank you to my beautiful sisters, Katie, Sarah and Chrissie, who have always been and always will be my best friends. Thank you to my Mom, who has provided endless encouragement and unconditional love in every single circumstance, and my Dad, who ignited my passion for science, nature, adventure, and the beauty of the oceans. A mere thank you does not suffice to express how grateful I am to my husband, Nick, for his unfailing support and love during these past few years. Thank you for being both my anchor to keep me grounded, and the wind in my sails to keep me going. Finally, my utmost thanks to God, who gave me a purpose. All my exploration into the nature of the ocean is an attempt to know You. This dissertation is dedicated to Aunt Leenie and Papa, who departed from this

Earth while I was on this graduate school journey. I love you and I miss you.

TABLE OF CONTENTS

LIST OF TABLES ...... ix LIST OF FIGURES ...... xi ABSTRACT ...... xiv

INTRODUCTION ...... 1

Chapter

1 CHEMICAL PREFERENCE AND ASSOCIATED BEHAVIORS OF FREE-SWIMMING SCLERACTINIAN CORAL LARVAE THROUGH EARLY DEVELOPMENT ...... 6

1.1 Abstract ...... 6 1.2 Introduction ...... 7 1.3 Methods ...... 10

1.3.1 Study site and organisms ...... 10 1.3.2 Larval collection and rearing ...... 11 1.3.3 Two-channel choice flume ...... 11 1.3.4 Still-water behavioral assay ...... 12 1.3.5 Chemical cues ...... 13 1.3.6 Video analysis ...... 13 1.3.7 Statistical analysis ...... 14

1.4 Results ...... 15

1.4.1 Acropora palmata ...... 17 1.4.2 Orbicella annularis ...... 23 1.4.3 Pseudodiploria strigosa ...... 25

1.5 Discussion ...... 27 1.6 Acknowledgements ...... 32

REFERENCES ...... 34

2 CORAL LARVAE BEHAVIORAL RESPONSE TO BENTHIC CUES DURING HABITAT SELECTION ...... 39

2.1 Abstract ...... 39 2.2 Introduction ...... 40 2.3 Methods ...... 43

2.3.1 Larval collection and rearing ...... 43 2.3.2 Cue generation ...... 43 2.3.3 Still-water behavioral assay ...... 45 2.3.4 Video analysis ...... 46 2.3.5 Statistics ...... 46

2.4 Results ...... 46

2.4.1 Brown algae ...... 46 2.4.2 Green algae ...... 49 2.4.3 Red algae ...... 50 2.4.4 Cyanobacteria ...... 50 2.4.5 Crustose coralline algae (CCA) ...... 50

2.5 Discussion ...... 51 2.6 Acknowledgements ...... 56

REFERENCES ...... 57

3 MULTIPLE ENVIRONMENTAL CUES IMPACT HABITAT CHOICE DURING NOCTURNAL HOMING OF SPECIALIZED REEF SHRIMP .... 63

3.1 Abstract ...... 63 3.2 Introduction ...... 64 3.3 Methods ...... 67

3.3.1 Temporal movements ...... 67 3.3.2 Natural habitat associations ...... 68 3.3.3 Morphological preference ...... 68 3.3.4 Chemical preference ...... 70 3.3.5 Patch reefs ...... 72

3.4 Results ...... 74

3.4.1 Temporal movements ...... 74 3.4.2 Natural habitat associations ...... 74 3.4.3 Morphological preference ...... 76 3.4.4 Chemical preference ...... 76 3.4.5 Patch reefs ...... 79

3.5 Discussion ...... 79 3.6 Acknowledgements ...... 83

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REFERENCES ...... 84

4 CORAL MORPHOLOGY IS LINKED TO INCREASED ABUNDANCE AND FITNESS OF OBLIGATE CORAL-DWELLING SHRIMP, CORALLIOCARIS GRAMINEA...... 88

4.1 Abstract ...... 88 4.2 Introduction ...... 89 4.3 Methods ...... 91 4.4 Results ...... 94 4.5 Discussion ...... 104 4.6 Acknowledgements ...... 107

REFERENCES ...... 108

5 CONCLUDING REMARKS ...... 112

REFERENCES ...... 114

Appendix

A SUPPLEMENTARY FIGURES ...... 115 B IMPACTS OF OCEAN ACIDIFICATION ON SENSORY PERCEPTION: UNDERLYING MECHANISMS AND TAXA COMPARISONS ...... 118

B.1 Introduction ...... 118 B.2 Chemosensory Perception ...... 120 B.3 Audition ...... 128 B.4 Vision ...... 132 B.5 Other senses ...... 135 B.6 Mechanisms ...... 137 B.7 Comparison between taxa ...... 139 B.8 Future directions ...... 141 B.9 Conclusions ...... 145 B.10 References ...... 145

C PERMISSIONS ...... 158

C.1 Multiple environmental cues impact habitat choice during nocturnal homing of specialized reef shrimp ...... 158 C.2 Impacts of Ocean Acidification on Sensory Function in Marine Organisms ...... 159

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LIST OF TABLES

Table 1.1: Number of days post fertilization during which behavioral assays could be conducted either in the two-channel choice flume or the still- water assay. The included n-values signify the final n-values used in the analyses after faulty videos were removed. N-values of O. annularis were low on day 6 and 7 due to limited number of larvae. Red X’s indicate days in which larvae of that species were not viable to be tested, either because they were not yet swimming (O. annularis, Day 3) or because they had already reached settlement-stage (all others)...... 16

Table 2.1: Functional and taxonomic groups of benthic algae and cyanobacteria used in this study (from 1Steneck and Dethier 1994, 2Littler et al. 1983, 3Diaz-Pulido 2010)...... 45

Table 3.1: Chemical choice comparisons. * indicates non-associated habitat based on transect data...... 72

Table 4.1: Observed and Expected shrimp abundance by coral species used in the χ2 goodness-of-fit test (χ2=32.442, p<0.001). The abundance of each coral species along the transect (column 2) was incorporated into the expected frequency of shrimp associations. Standardized Residuals (Std. Res.) indicate the extent and direction of deviation from expected values...... 95

Table 4.2: Negative binomial generalized linear model to determine influence of coral morphology on quantity of C. graminea...... 96

Table 4.3: Multiple linear regression results to predict body size characteristics (TL, SL, carapace width). Diameter, height and sex were used as predictors in the model. The only significant interaction effect was between coral diameter and shrimp sex. Regression graphs are depicted in Appendix A. a=0.05, * indicates significance...... 100

Table 4.4: Multiple regression analysis to determine body size characteristics (TL, SL, carapace width), using inter-brachial spacing and sex as predictors. The interaction between sex and inter-brachial spacing was significant and remained in the model. a=0.05, * indicates significance...... 100

Table 4.5: Multiple linear regression results to predict shrimp claw dimensions (length, width, L/W ratio). Diameter, height and sex were used as predictors in the model. The only significant interaction effect was between coral diameter and shrimp sex. Regression graphs are depicted in Appendix A. a=0.05, * indicates significance...... 101

Table 4.6: Multiple regression analysis to determine shrimp claw dimensions (length, width, L/W ratio), using inter-brachial spacing and sex as predictors. The interaction term was not significant and was thus removed from the model. a=0.05, * indicates significance...... 101

Table 4.7: Multiple regression analysis to determine influence of body (TL, SL, carapace width) and claw (length, width, ratio) dimensions on shrimp fecundity measured by egg count. Body size predictors and claw size predictors were analyzed in two separate models. No interaction terms were significant were thus removed from the model. a=0.05, * indicates significance ...... 103

Table 4.8: Simple linear regressions to determine correlation between C. graminea fitness (body size, claw size, fecundity) and the number of resident shrimp within each coral colony. a=0.007 after Bonferroni correction, * indicates significance...... 103

Table B.1: Synopsis of research studies evaluating the influence of ocean acidification on chemosensory perception. The table highlights focal species, experimental CO2 levels, control CO2 levels, sensitivity at which an effect was observed and methodology. NH = Newly Hatched, * indicates species was treated with high CO2 starting at birth/hatching...... 122

Table B.2: Synopsis of research studies evaluating the influence of ocean acidification on auditory perception and otolith development. The table highlights focal species, experimental CO2 levels, control CO2 levels, sensitivity at which an effect was observed and methodology. NH = Newly Hatched, * indicates species was treated with high CO2 starting at birth/hatching...... 130

Table B.3: Synopsis of research studies evaluating the influence of ocean acidification on visual perception. The table highlights focal species, experimental CO2 levels, control CO2 levels, sensitivity at which an effect was observed, methodology and the specific component of vision that was tested. NH = Newly Hatched, * indicates species was treated with high CO2 starting at birth/hatching...... 133

LIST OF FIGURES

Figure 1.1: Chemical preferences of Acropora palmata larvae throughout early development in the two-channel choice flume. Mean percent time (±SE) spent in general reef water (dark blue bars) or offshore water (light blue bars) is depicted for each day that the larvae were free- swimming. Wilcoxon signed rank test, a=0.05, * indicates significance...... 17

Figure 1.2: Variations in mean percent time (±SE) spent traveling during early development of (A) Acropora palmata, (B) Orbicella annularis, and (C) Pseudodiploria strigosa larvae after they were presented with either general reef water (dark blue bars) or offshore water (light blue bars) ANCOVA, a=0.05, * indicates significance...... 19

Figure 1.3: Variations in mean velocity (±SE) during early development of (A) Acropora palmata, (B) Orbicella annularis, and (C) Pseudodiploria strigosa larvae when presented with either general reef water (dark blue bars) or offshore water (light blue bars). ANCOVA, a=0.05, * indicates significance...... 20

Figure 1.4: Variations in behavior of Acropora palmata larvae during early development. The mean percent time (±SE) spent displaying each of the four behavioral classifications is depicted for each day (listed at the top) and each cue (listed at the bottom). ANCOVA, a=0.05, * indicates significance...... 22

Figure 1.5: Variations in behavior of Orbicella annularis larvae during early development. The mean percent time (±SE) spent displaying each of the four behavioral classifications is depicted for each day (listed at the top) and each cue (listed at the bottom). ANCOVA, a=0.05, * indicates significance...... 24

Figure 1.6: Variations in behavior of Pseudodiploria strigosa larvae during early development. The mean percent time (±SE) spent displaying each of the four behavioral classifications is depicted for each day (listed at the top) and each cue (listed at the bottom). ANCOVA, a=0.05, * indicates significance...... 26

Figure 2.1: Comparison of mean percent time (±SE) spent traveling before (lighter colors) versus after (darker colors) the chemical cue of each benthic species was inserted into the petri dish. Brown bars=brown algae, green bars=green algae, red bars=upright red algae, blue bars= cyanobacteria, purple bars=CCA. Wilcoxon signed rank test, a=0.05, * indicates significance...... 47

Figure 2.2: Comparison of mean velocity (±SE) before (lighter colors) versus after (darker colors) the chemical cue of each benthic species was inserted into the petri dish. Brown bars=brown algae, green bars=green algae, red bars=upright red algae, blue bars= cyanobacteria, purple bars=CCA. Wilcoxon signed rank test, a=0.05, * indicates significance...... 48

Figure 2.3: Comparison of pre-cue and post-cue mean percent time (±SE) spent in each of the four behavioral classifications, (A) Stationary/Rotations, (B) Small Circles, (C) Non-Directed Movement, and (D) Directed Movement. Lighter colors refer to pre-cue data and darker colors refer to post-cue data. Brown bars=brown algae, green bars=green algae, red bars=upright red algae, blue bars= cyanobacteria, purple bars=CCA. Wilcoxon signed rank test, a=0.05, * indicates significance...... 49

Figure 3.1: Infogram of the experimental methods to test sponge morphology preferences in isolation. Grey cylinders represent artificial sponges made of foam pool noodles. Experimental results are shown in text boxes with a significant preference for tall and narrow tube morphology...... 70

Figure 3.2: Infogram of the experimental methods to test sponge preference when all cues were present, and sponges were equidistant. Experimental results are shown in text boxes with a significant preference for C. vaginalis...... 73

Figure 3.3: Natural shrimp-sponge associations. Transect data reveals a preference for tall narrow sponge tubes when all sponges are combined and when comparing C. vaginalis alone. White bars represent sponge tubes that did not host shrimp while grey bars represent sponge tubes with resident shrimp. * indicates significance. .. 76

Figure 3.4: Chemical choice trials. (A) Shrimp response to individual sponge cues compared to general reef cues. (B) Shrimp response when presented with two sponge cues simultaneously to determine a hierarchical preference. All chemical data are mean percent time ± standard error. * indicates significance...... 78

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Figure 4.1: Focal shrimp, Coralliocaris graminea, depicting body (Standard length (SL), Total length (TL), Carapace width) and claw (length, width) measurements. Inlay: Posterior ventral view of gravid female C. graminea...... 93

Figure 4.2: Habitation of C. graminea within Acropora corals is associated with (A) coral head diameter and (B) coral branch height, but not with (C) inter-branchial spacing. Dark grey bars represent corals without resident C. graminea while light grey bars represent corals with shrimp inhabitants. Wilcoxon signed rank test, a=0.05, * indicates significance...... 96

Figure 4.3: Sex ratios of C. graminea within Acropora corals. 69 shrimp were collected from 31 coral individuals: 44 females, 18 males, and 7 juveniles...... 97

Figure 4.4: Sexual dimorphism between male and female C. graminea. Males are represented by the dark grey bars while females are represented by the light grey bars. t-test, a=0.05, * indicates significance...... 98

Figure A.1: Linear regression graphs displaying relationship between body size characteristics (TL, SL, carapace width; y-axis) and coral morphology (diameter, height; x-axis). Female shrimp are depicted with red markers while males are depicted with blue markers. Full statistical outputs are described in Figure 4.3...... 115

Figure A.2: Linear regression graphs displaying relationship between claw size characteristics (length, width, y-axis) and coral morphology (diameter, height; x-axis). Female shrimp are depicted with red markers while males are depicted with blue markers. Full statistical outputs are described in Figure 4.5...... 116

Figure A.3: Linear regression graphs displaying relationship between fecundity (number of eggs) and coral morphology (diameter, height; x-axis). Full statistical outputs are described in Figure 4.5...... 117

Figure B.1: Infogram depicting the summarized impacts of ocean acidification on the sensory systems of a model vertebrates and invertebrate organisms...... 142

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ABSTRACT

Habitat selection is a critical process for most organisms that are mobile for at least a portion of their lives and can influence survival and lifelong fitness. Selection of optimal habitat is accomplished through the detection and assessment of environmental cues such as waterborne chemicals or structural morphology. Substrate availability on coral reefs is limited, giving rise to high competition between benthic species. Here, I assessed different aspects of sensory-mediated habitat selection of reef invertebrates through a variety of research methods, invertebrate species, and geographic locations to develop an integrated and diverse understanding of the process and implications of habitat selection. First, habitat selection is not a singular event, but rather, an evolving process throughout development. I investigated the developmental variations in coral larvae preference and associated behaviors in response to general reef cues using three species of Caribbean corals. Larvae of Acropora palmata, Orbicella annularis, and Pseudodiplora strigosa were exposed to general reef or offshore cues each day of their larval duration. A. palmata were tested in both flowing water, using a two-channel choice flume, and in a novel still-water behavioral assay; due to weaker swimming capabilities, the other species were only tested in the still-water assay. A. palmata larvae prefer reef cues compared to offshore cues throughout development, but all three species displayed minute, yet distinct, behaviors that varied based on cue and developmental stage. These behaviors may be indicative of emerging sensitivities to environmental stimuli and should be investigated further.

xiv

Sensory-mediated habitat selection is not solely dependent on general reef cues but may also rely on specific cues of the benthic reef environment. A. palmata were tested in the still-water behavioral assay to determine attraction or repulsion to algal and cyanobacterial chemical cues. Regardless of benthic species, coral larvae increased the percent of time spent moving after the cues were presented. Notably, coral larvae significantly increased the proportion of time spent in directed motion or swimming in small circles only when presented with crustose coralline algae (CCA), potentially revealing these minute behaviors as indicators of positive habitat cues. Habitat selection is not only relevant early in organismal life history, but is also vital for roving, migrating, or homing adults. Sensory-mediated adult habitat selection was investigated in the specialized reef shrimp, Lysmata pederseni, which resides within Caribbean tube sponges, but nocturnally vacates to feed or mate. Using mark and recapture techniques, reef transects, cue isolation experiments, and patch reefs, I discovered that while shrimp use both chemical and morphological cues to assess potential habitats, the optimal sponge habitat is defined more by shape than by chemistry. Morphological characteristics of specialized habitat are further investigated in my final chapter, where I attempt to relate habitat to fitness of a coral-dwelling Indo-

Pacific shrimp, Coralliocaris graminea. Transects reveal non-random associations of shrimp within 7 species of Acropora corals. Shrimp abundance is closely related to coral morphology, especially coral head diameter and branch height. Further, C. graminea individuals were collected from host corals to reveal that many aspects of body and claw size, as well as fecundity, can be positively correlated with coral size.

The potential exists for this habitat-specialized shrimp to survive within structurally similar, non-host corals if preferred hosts are stressed or degrading. In all, this body of work contributes to our understanding of habitat selection, from ontogenesis to adults. Knowledge of both the environmental stimuli that influence habitat selection and the implications of that choice can guide research and management practices. An integrated approach was taken to each of these projects, where multiple methods were incorporated and compared to determine varying behaviors, motivations, and repercussions of correctly interpreting stimuli to select the optimal habitat. However, during the current period of unprecedented environmental change, the optimal habitats are changing and so are the various cues by which organisms can detect and select this habitat. This research has made strides towards not only understanding habitat selection of cryptic, specialized, and/or foundational species, but also predicting how these sensory-mediated behavioral responses might shift as environmental conditions decline.

xvi

INTRODUCTION

Habitat selection in the marine environment is a function of physical processes and animal behavior (Meadows and Campbell 1972; Cowan and Sponaugle 2009).

The relative influence of each of these factors depends on size, swimming capabilities, responsiveness to cues, and necessity of specific habitats for individual species

(Meadows and Campbell 1972; Pawlik 1992; Raimondi and Morse 2000). Habitat selection occurs at several points throughout organismal life history (Gillanders et al.

2003), from initial habitat selection early in development (Buttman 1987; Lecchini et al. 2005) to homing or migration of adults (Thorrold et al. 2001; White and Brown

2013). In highly complex coral reefs, available substrate is limited; while appropriate habitat is important for all organisms, it is especially critical for animals that require specialized habitat (Wilson et al. 2008) or are permanently sessile once habitat has been selected (Hadfield 1986). In both situations, the selection of inappropriate habitat can lead to reduced fitness or, in some cases, death (Raimondi and Morse 2000).

Organisms orient to and choose between habitats using available environmental cues, including waterborne chemicals, soundscapes, and structural morphology, assessed through visual and tangible cues. Chemosensory reception is one of the primary senses used by most marine organisms to detect habitat (Atema et al. 2002; Hay 2009) and chemical cues can be detected up to 2km from the source

(Lecchini et al. 2014). The chemical signature of a coral reef relays information

1 regarding the health of the ecosystem (Lecchini et al. 2013; Dixson et al. 2014).

Additionally, soundscapes made of snapping shrimp and fish vocalizations allow potential recruits to successfully orient to coral reefs (Simpson et al. 2005). Visual and tactile cues can inform organisms about the morphological characteristics of potential habitats to ensure the optimal location is chosen (Ambrosio and Brooks 2011).

Coral reefs around the world are experiencing unprecedented deterioration

(Hughes 1994; Hoegh-Guldberg et al. 2007). Environmental stressors from myriad sources contribute to phase shifts from coral to algal dominated reefs (Done 1992;

Knowlton 1992), with associated reductions in biodiversity and species abundance.

Consequently, the environmental cues that organisms use to select habitat reflect these changes. For example, chemical signatures of reefs shift as species compositions change; dead and living reefs not only produced different chemicals, but these chemical differences were detectable to reef fish, with fish preferring the cues associated with a living reef (Lecchini et al. 2014).

This compilation of studies investigates habitat selection of reef invertebrates at different life stages. First, the development of chemical sensitivity to the reef environment is assessed in three species of Caribbean coral larvae (chapter 1).

Subsequently, coral larval response to specific components of the reef were observed to understand the influence of benthic cues on habitat selection of reef-building corals

(chapter 2). From there, I investigated which environmental cues - chemical or morphological - were more relevant for habitat selection of an adult Caribbean reef shrimp (chapter 3). Finally, habitat associations of an Indo-Pacific reef shrimp were

2 analyzed in an attempt to correlate habitat with organismal fitness, a potential long- term consequence of habitat selection (chapter 4).

REFERENCES

Alvarez-Filip L, Dulvy NK, Gill JA, Côté IM, Watkinson AR. 2009. Flattening of Caribbean coral reefs: region-wide declines in architectural complexity. Proc R Soc Lond B Biol Sci. 276:3019-3025.

Ambrosio LJ, Brooks WR. 2011. Recognition and use of ascidian hosts, and mate acquisition by the symbiotic pea crab Tunicotheres moseri (Rathbun, 1918): the role of chemical, visual and tactile cues. Symbiosis. 53:53-61.

Atema J, Kingsford MJ, Gerlach G. 2002. Larval reef fish could use odour for detection, retention and orientation to reefs. Mar Ecol Prog Ser. 241:151-160.

Butman CA. 1987. Larval settlement of soft-sediment invertebrates: the spatial scales of pattern explained by active habitat selection and the emerging role of hydrodynamical processes. Oceanogr Mar Biol. 25:113-165.

Cowen RK, Sponaugle S. 2009. Larval dispersal and marine population connectivity. Ann Rev Mar Sci. 1:443-466.

Dixson DL, Abrego D, Hay ME. 2014. Chemically mediated behavior of recruiting corals and fishes: a tipping point that may limit reef recovery. Science. 345:892-897.

Done TJ. 1992. Phase shifts in coral reef communities and their ecological significance. Hydrobiologia. 247:121-32.

Gillanders BM, Able KW, Brown JA, Eggleston DB, Sheridan PF. 2003. Evidence of connectivity between juvenile and adult habitats for mobile marine fauna: an important component of nurseries. Mar Ecol Prog Ser. 247:281-295.

Hadfield MG. 1986. Settlement and recruitment of marine invertebrates: a perspective and some proposals. Bull Mar Sci. 39:418-25.

Hughes TP. 1994. Catastrophes, phase shifts, and large-scale degradation of a Caribbean coral reef. Science. 265:1547-51.

Hay ME. 2009. Marine chemical ecology: chemical signals and cues structure marine populations, communities, and ecosystems. Ann Rev Mar Sci 1:193-212

Hoegh-Guldberg O, Mumby PJ, Hooten AJ, Steneck RS, Greenfield P, Gomez E, Harvell CD, Sale PF, Edwards AJ, Caldeira K, Knowlton N. 2007. Coral reefs under rapid climate change and ocean acidification. Science. 318:1737-1742.

Knowlton, N. 1992. Thresholds and multiple stable states in coral reef community dynamics. Am Zool. 32:674–682.

Lecchini D, Shima J, Banaigs B, Galzin R. 2005. Larval sensory abilities and mechanisms of habitat selection of a coral reef fish during settlement. Oecologia. 143:326-34.

Lecchini D, Waqalevu VP, Parmentier E, Radford CA, Banaigs B. 2013. Fish larvae prefer coral over algal water cues: implications of coral reef degradation. Mar Ecol Prog Ser. 475:303-307.

Lecchini D, Miura T, Lecellier G, Banaigs B, Nakamura Y. 2014. Transmission distance of chemical cues from coral habitats: implications for marine larval settlement in context of reef degradation. Mar Biol. 161:1677-86.

Meadows PS, Campbell JI. 1972. Habitat selection by aquatic invertebrates. Adv Mar Biol. 10:271-382.

Pawlik JR. 1992. Chemical ecology of the settlement of benthic marine invertebrates. Oceanogr Mar Biol Annu Rev. 30:273-335.

Raimondi PT, Morse, A.N. 2000. The consequences of complex larval behavior in a coral. Ecology. 81:3193-3211.

Simpson SD, Meekan M, Montgomery J, McCauley R, Jeffs A. 2005. Homeward sound. Science. 308:221.

Thorrold SR, Latkoczy C, Swart PK, Jones CM. 2001. Natal homing in a marine fish metapopulation. Science. 291:297-9.

White GE, Brown C. 2013. Site fidelity and homing behaviour in intertidal fishes. Mar Biol. 160:1365-72.

Wilson SK, Burgess SC, Cheal AJ, Emslie M, Fisher R, Miller I, Polunin NV, Sweatman HP. 2008. Habitat utilization by coral reef fish: implications for specialists vs. generalists in a changing environment. J Anim Ecol. 77:220- 228.

Chapter 1

CHEMICAL PREFERENCE AND ASSOCIATED BEHAVIORS OF FREE- SWIMMING SCLERACTINIAN CORAL LARVAE THROUGH EARLY DEVELOPMENT

1.1 Abstract

Most coral species have a split life cycle, where a relatively sessile adult produces free-swimming larvae or planula. Larvae of broadcast-spawning coral species are subject to physical ocean processes and may develop offshore before returning to the reef to settle. Orienting back to the reef environment requires the use of numerous environmental cues, including water-soluble chemicals. Sensitivity and attraction to these cues may change as larvae develop and seek settlement sites. Temporal variations in coral larvae behavior when exposed to general reef cues was investigated using three species of Caribbean coral. Larvae of Acropora palmata, Orbicella annularis, and Pseudodiploria strigosa were collected in situ and reared until they were capable of swimming before being exposed to reef and offshore cues for two separate assays: a two-channel choice flume and a novel still-water behavioral assay. The choice flume experiment revealed that A. palmata invariably preferred the chemical cues associated with a reef compared to offshore cues. However, in the still- water assay larvae displayed a reversal in some behaviors based on cue depending on developmental stage; the proportion of time coral larvae spent traveling through the dish was greater in offshore water early in development, but greater in reef water later in development. For all three coral species, four distinct behaviors were characterized and recorded, with reef cue initiating significant changes in time spent stationary or in

6 non-directional movement. These behaviors may be analogous to benthic probing or testing behaviors and potentially indicate recognition of preferred habitat cues. Investigations of chemical preferences and the associated minute behaviors of coral larvae throughout development can augment our understanding of the process of habitat selection in foundational coral species.

1.2 Introduction Coral reefs have been described as the rainforests of the ocean, accounting for more than a quarter of all marine species (Knowlton et al. 2010). Like the trees in rainforests, coral colonies create the architecture of the ecosystem and provide dozens of ecological services to the local denizens (Moberg and Folke 1999). Unlike trees, however, corals can actively select their habitat during a self-locomotive larval stage (Raimondi and Morse 2000). In the marine environment, most organisms that are permanently sessile at the adult stage have a motile larval stage that allows for behavioral choices before and during the settlement process (Chia 1989; Pawlik 1992; Rodríguez et al. 1993; Katz et al. 1994; Tamburri et al. 1996; Raimondi and Morse 2000; Jenkins 2005). These choices are mediated by environmental cues perceived by the sensory systems of the individual (Hadfield and Paul 2001; Vermeij et al. 2010). For example, barnacle larvae use the chemical cues of conspecifics to navigate to appropriate sites for gregarious settlement (Knight-Jones 1953; Burke 1986). An appropriate settlement site among barnacle conspecifics increases fitness because proximity facilitates breeding (Buschbaum 2001). Corals reproduce by either brooding or broadcast spawning (Szmant 1986). Brooders produce fully-functional planulae that settle immediately, often in the vicinity of their parental colony. Broadcast spawners, however, release gametes into

7 the water column for fertilization, a process that typically occurs annually. The fertilized cells develop in the water column and are often transported by currents from their native reef (Wood et al. 2014). Coral larvae have a limited free-swimming period before settlement, and choosing an appropriate settlement site is critical, as nascent corals are permanently sessile after metamorphosis (Burke 1983). Motile coral larvae must first select habitat at the ecosystem level using appropriate whole-reef cues and then must find suitable settlement sites within the reef matrix using specific benthic cues (Ritson-Williams et al. 2010, 2016). This process is mediated by environmental cues such as light (Maida et al. 1994; Mundy and Babcock 1998), sound (Vermeij et al. 2010), pressure (Stake and Sammarco 2003), and soluble chemicals of both the general reef and specific benthic species (Dixson et al. 2014). Research into chemically-mediated habitat selection in corals has primarily focused on the near-settlement stage larvae that are actively probing the bottom (Hadfield and Paul 2001; Gleason et al. 2009; Price 2010; Tebben et al. 2015). Laboratory and field-based settlement assays help to determine the chemical cues involved in both settlement and metamorphosis. Using a two-channel choice flume, Dixson et al (2014) found that pre-settlement stage, free-swimming coral larvae are capable of distinguishing chemical signatures of both the general reef and specific benthic components. Indo-Pacific acroporid species swam towards the chemical cues of healthy, diverse reefs yet actively avoided the chemical cues of degraded, algae- dominated reefs. The next step in this field is to determine if this behavior is universal to all coral species and across ocean ecosystems. Further, the timing of these behaviors will augment our understanding of the life history of these vital reef builders. For example,

8 during pelagic development anemonefish display alternating chemical preferences for their host sea anemones compared to untreated water; larval anemonefish avoid water containing habitat-rich cues until mid-way through their developmental pelagic period when returning to the reef for settlement is required (Dixson et al. 2011). It is unknown if coral larvae share this fluctuating preference. During the planktonic development of scleractinian coral larvae, lipid energy reserves are depleted (Arai et al. 1993; Harii et al. 2007), locomotive cilia form, and epidermal cells specialize into sensory and secretory cells (Heyward 1987). As settlement competency approaches, larvae become increasingly mobile and more sensitive to external cues. Sensory cells are concentrated at the aboral end, from which they typically probe the substrate and eventually attach (Tran and Hadfield 2013). Sensory capabilities and associated behaviors during this critical planktonic period between fertilization and settlement can impact individual survival and population replenishment. Due to species differences in larval size and swimming capabilities, the two- channel choice flume used in previous experiments may not be an appropriate behavioral assay for all coral species, as it requires the larvae to continually swim against a 70 mL min-1 (0.72 cm sec-1) flow rate. Additionally, the flume protocol can only determine chemical preferences but cannot assess minute changes in behavior associated with environmental cues, such as changes in velocity. To address these issues, I created a novel assay to test behavioral changes to environmental cues for coral species that are not compatible in the flume. This chapter investigates both the chemical preferences and the associated behaviors of coral larvae throughout their free-swimming development. By video recording still-water assays after reef cues have been introduced to three species of

9 Caribbean coral larvae, I investigated the influence of early development on cue sensitivity and associated behaviors. Behavioral data are viewed in light of data collected using the traditional flume method to determine preference or avoidance of general reef and offshore cues. I hypothesized that coral larvae would exhibit differential behaviors depending on the cue that was presented and that these behaviors would shift as the larvae approached settlement. This research allows us to compare species-specific developmental shifts in response to reef chemistry in order to gain a more thorough understanding of how coral larvae select habitat. This is especially critical for coral species that are endangered, such as two of the focal species, as knowledge of habitat preferences can inform conservation practices.

1.3 Methods

1.3.1 Study site and organisms

Research was conducted at the Smithsonian Carrie Bow Cay field station off the coast of Belize (16°48'9.26"N, 88° 4'54.87"W). The larvae of three Caribbean scleractinian, corals were used in the behavioral assays. Acropora palmata is a foundational branching coral among Caribbean reefs but is critically endangered and has faced as much as 80% reduction in coverage (Bruckner 2002; National Marine

Fisheries Service 2006). Orbicella annularis (previously Montastraea annularis) and Pseudodiploria strigosa (previously Diploria strigosa) are both massive boulder corals dominant in the Caribbean, however, O. annularis has been classified as endangered due to excessive population declines (Edmunds and Elahi 2007; National Marine Fisheries Service 2014). All three species are broadcast spawners and usually experience a peak spawn in August. Typically, A. palmata synchronously spawns 3-5

10 days after the full moon and up to 4.5 hours after sunset (Jordan 2018). O. annularis and P. strigosa spawn later in both regards – usually 6-8 days after the full moon and up to 5.5 hours after sunset (Jordan 2018). Larvae of all three species are lecithotrophic and depend on lipid-rich energy reserves during their planktonic period (Baird et al. 2009).

1.3.2 Larval collection and rearing Acropora palmata larvae were collected on 17 August 2014 and 5-6 August 2015, while Orbicella annularis and Pseudodiploria strigosa larvae were collected on 8 August 2015. Coral colonies (<4m depth) were observed by divers and tented similar to Sharp et al. (2010), once pre-released gamete bundles were observed at the tip of the polyps. Upon spawning, the positively buoyant gamete bundles accumulated in a falcon tube at the top of the tent structure. After spawning, bundles were transported to the Carrie Bow Cay field station for fertilization. Eggs and sperm from separate colonies were cross-fertilized for one hour before the excess sperm was rinsed off with seawater. Coral larvae were reared in 4L larval containers with continual seawater flow (Ritson-Williams et al. 2010). Environmental conditions of the rearing tanks, i.e. temperature and light cycle, mirrored conditions found in the natural reef environment. Throughout development, deceased larvae and debris were removed from the containers with pipettes.

1.3.3 Two-channel choice flume

A two-channel choice flume (Gerlach et al. 2007) was used to assess chemical preference of coral larvae to either offshore or general reef cues throughout early development. The choice flume (23.5 x 4 cm) allows an individual organism to

11 experience the chemical cues of two different water sources simultaneously by presenting them side-by-side. Water from the two sources were gravity-fed from buckets through tubing and into the choice chamber. Laminar flow was maintained at 70 mL min-1 (0.72 cm sec-1) using flow meters (Dwyer MMA-40) and checked periodically using dye tests. A coral larva was pipetted into the center of the flume and given a 30s habituation period, during which time the larva was free to explore either cue. During the habituation period, individuals were carefully observed to ensure that they swam against the current. Individuals that could not maintain their position were removed from the trial because they likely indicated a lack of swimming capability rather than a lack of preference. After the habituation period, larva position in each stimulus was recorded at 5s intervals for 2 min. The larva was then removed from the flume for 1 minute while the water sources were switched to eliminate a potential side bias. The same individual was then placed back into the center of the flume for another 30s habituation and 2 min test. Experimenters were unaware of which side held which cue and thus remained blind during the entirety of the trials to avoid bias.

1.3.4 Still-water behavioral assay Plastic petri dishes (90 mm) were marked with a 1 cm2 grid and filled with 30 mL filtered seawater. Microscope lights (Omano, MikeLite) with duel gooseneck arms illuminated the dish from opposite sides such that the photosynthetically active radiation (PAR) at the surface of the dish was 95-111 µmol m-2 s-1 (Hansatech light meter). Cameras (Apple iPod Touch) attached to adjustable tripods were positioned above the petri dishes for aerial footage of each behavioral assay. An individual coral larva was pipetted into the center of the petri dish and allowed a 1 min habituation period, after which video recording commenced. After 2 min, the larva was relocated

12 back to the center of the dish using a glass pipette. Immediately, 2 mL from each water source was carefully inserted into opposite sides of the dish using 5 mL syringes. Video recording continued for 2 min before the conclusion of the trial. The cue was inserted into alternating sides of the dish for each trial to avoid any potential side bias. Dye tests revealed the chemical plume distance to be 3-4 cm from the insertion point.

1.3.5 Chemical cues During both the choice flume and the still-water assay, larvae were presented with reef water and offshore water each day (n=20 for each assay). Individual larvae were only used for one trial. An additional 20 larvae per assay per day acted as a control and were tested when exposed to offshore water on both sides. Reef water was collected by boat from surface water flowing over a reef <3 m below the surface. Offshore water was collected by boat >1 km from the nearest land mass. Water cues were used in behavioral assays within 6 hours after collection.

1.3.6 Video analysis

Behavioral videos were analyzed using Tracker 5.0 (Brown 2019) to assess velocity. Autotracker was initiated when conditions permitted (i.e. the larva was distinguishable from the background), and larval position was tracked manually when autotracking failed. Video frame rate was 29.97 frames/s and larval position was recorded every 5 frames. Frames involving larval relocation or cue insertion were removed from the analysis. Behavioral videos were also analyzed manually to characterize behavior type. Four general behaviors were identified: (1) Stationary Rotations, in which the larva either remained motionless or rotated in place, (2) Small

13 Circles, in which the larva swam in confined circles <1 cm, yet did not experience any overall change in position within the dish, (3) Directed Movement, in which the larva swam in one direction with limited variations in speed; and (4) Non-directed Movement, in which the larva traveled in an unpredictable fashion involving spiraling, drifting, bouncing, or serpentine motion. For both Tracker and manual analyses, the first 10 seconds of footage after the cue was introduced was removed to account for any water movement due to the induction of the cue rather than larval self-propulsion.

1.3.7 Statistical analysis Chemical choice fluming experiments were analyzed using the Wilcoxon signed rank test (JMP Pro 13). For the still-water analysis, average velocity and percent time spent in each behavior were analyzed in R (R Core Team 2017) using a between subjects analysis of covariance (ANCOVA) to assess the effect of cue type, after controlling for pre-cue insertion data. Post-cue data (average velocity or percent time in specific behaviors after the cue was presented) acted as the dependent variable, cue type (reef vs offshore) acted as the independent variable, and pre-cue data (average velocity or percent time in specific behaviors before the cue was presented) acted as the covariate. Data from each day post fertilization were analyzed separately because the covariate, pre-cue data, can be influenced by developmental age. In each test, the data was screened for assumptions and tested for outliers using Mahalanobis distance. When an outlier was detected (n=2), it was removed from the analysis. Linearity, homogeneity (Levene’s tests), homoscedasticity, and normality (Shapiro- Wilks test) were verified for all tests. When data were not normally distributed, a square-root transformation was applied.

14 1.4 Results Larvae of A. palmata, O. annularis, and P. strigosa had varied developmental times and swimming capabilities. For the fluming trials, larvae were required to swim against a 70mL min-1 flow rate. This swimming speed was attainable for A. palmata larvae on developmental days 4-6 (Table 1.1). Larvae of O. annularis and P. strigosa were never capable of swimming against this current and were thus incapable of being tested in the two-channel choice flume. In contrast, the still-water behavioral assays were an appropriate method of observing larval behavior throughout development, regardless of swimming capabilities. Experiments were conducted on each species during their early development only while swimming was detectable, i.e. beginning when larvae first showed signs of movement and ending when larvae immediately settled when placed in the assay and did not react to any cue (Table 1.1). O. annularis larvae were actively swimming on day 7 post-fertilization; however, a population decline in the reared larvae forced an early termination of the experiment for that species, as well as a reduced number of individuals per test on days 6 (n=16) and 7 (n=10). A fraction of videos could not be analyzed and were removed from the analysis. Final n-values of each trial are provided in Table 1.1.

15 Table 1.1: Number of days post fertilization during which behavioral assays could be conducted either in the two-channel choice flume or the still-water assay. The included n-values signify the final n-values used in the analyses after faulty videos were removed. N-values of O. annularis were low on day 6 and 7 due to limited number of larvae. Red X’s indicate days in which larvae of that species were not viable to be tested, either because they were not yet swimming (O. annularis, Day 3) or because they had already reached settlement-stage (all others).

Day 3 Day 4 Day 5 Day 6 Day 7

Choice Choice Choice

flume: n=20 flume: n=20 flume: n=20 Still-water

assay: Still-water Still-water Still-water Reef: n=20 assay: assay: assay: X palmata

Offshore: . Reef: n=18 Reef: n=20 Reef: n=19 A n=18 Offshore: Offshore: Offshore: n=20 n=20 n=19

Still-water Still-water Still-water Still-water assay: assay: assay: assay: X Reef: n=18 Reef: n=18 Reef: n=13 Reef: n=8 annularis Offshore: Offshore: Offshore: Offshore:

O n=16 n=17 n=15 n=9

Still-water Still-water assay: assay: Reef: n=17 Reef: n=17

strigosa X X X Offshore: Offshore:

P. n=19 n=19

16 1.4.1 Acropora palmata A. palmata larvae discriminated reef and offshore chemical cues within the two-channel choice flume between days 4-6 post-spawn. Larvae consistently spent more time in the flume side containing cues associated with reef water compared to offshore water (p<0.001 for all trials) (Figure 1.1).

Figure 1.1: Chemical preferences of Acropora palmata larvae throughout early development in the two-channel choice flume. Mean percent time (±SE) spent in general reef water (dark blue bars) or offshore water (light blue bars) is depicted for each day that the larvae were free-swimming. Wilcoxon signed rank test, a=0.05, * indicates significance.

17 Even though larvae could not swim against the flume current earlier than day 4, behavioral changes could be observed in the still-channel assay on days 3-6. The percentage of time spent traveling was divergent between cue types for A. palmata larvae (Figure 1.2A). On day 3, larvae spent significantly more time traveling only when exposed to the offshore cues (p=0.022), whereas on day 5 and 6, larvae spent significantly more time traveling when the reef cue was added (p=0.022, p=0.028, respectively). Average velocity was higher on day 3 and 4 when larvae were presented with only the offshore cue (p<0.001, p=0.026, respectively) (Figure 1.3A). However, on day 5 and 6, no difference in velocity was detected.

19 A. Acropora palmata 0.2 *

0.15 *

0.1

0.05

Day 3 Day 4 Day 5 Day 6 B. Orbicella annularis 0.15

0.1

0.05 Mean Velocity (cm/s) Velocity Mean

Day 4 Day 5 Day 6 Day 7 C. Pseudodiploria strigosa 0.15 Reef H2O

Offshore H2O 0.1

0.05

Day 3 Day 4 Figure 1.3: Variations in mean velocity (±SE) during early development of (A) Acropora palmata, (B) Orbicella annularis, and (C) Pseudodiploria strigosa larvae when presented with either general reef water (dark blue bars) or offshore water (light blue bars). ANCOVA, a=0.05, * indicates significance.

20 Four broad behavioral classifications were identified during the video analysis and each were analyzed separately (Figure 1.4). On day 3, A. palmata larvae spent a greater amount of time moving in a directed manner when presented with the offshore cue (p=0.021). On day 4, larvae spent more time stationary or performing rotations in place when presented with the reef cue (p=0.025). On day 5 and 6, the percent time spent on each of the four behaviors did not significantly vary between the cues. Small circles were a distinct, but relatively rare behavior regardless of the day or cue and could not be analyzed with this method as the distributions were not normal, despite transformations.

21 Acropora palmata Day 3 Day 4 Day 5 Day 6 50

* Directed Movement 40

Non-directed Movement 50

Small Circles 30

% time spent displaying each behavior each displaying spent time % 50

* Stationary/Rotations 40

0 Reef Offshore Reef Offshore Reef Offshore Reef Offshore Figure 1.4: Variations in behavior of Acropora palmata larvae during early development. The mean percent time (±SE) spent displaying each of the four behavioral classifications is depicted for each day (listed at the top) and each cue (listed at the bottom). ANCOVA, a=0.05, * indicates significance.

22 1.4.2 Orbicella annularis O. annularis larvae did not significantly vary in the amount of time spent travelling on day 4, 5 or 7. On day 6, however, larvae traveled a greater percentage of time when exposed to offshore cues (p=0.049) (Figure 1.2B). Despite variations in travel time, O. annularis larvae did not experience any changes in velocity in response to cue type, regardless of developmental day (Figure 1.3B). Behavior type varied significantly on day 4 and 5 (Figure 1.5). On day 4, larvae spent more time stationary while in offshore water compared to reef water (p=0.031). This behavior shifted on day 5, with larvae spending more time stationary when presented with reef water (p=0.038). This increase in time spent stationary in reef water was paired with a significant reduction in time spent in non-directed movement (p=0.021).

23 Orbicella annularis

40 Day 4 Day 5 Day 6 Day 7 Directed Movement

0 70 *

60 Non-directed Movement

Small Circles 30

% time spent displaying each behavior each displaying spent time % 10

Stationary/Rotations 40 * * 30

0 Reef Offshore Reef Offshore Reef Offshore Reef Offshore Figure 1.5: Variations in behavior of Orbicella annularis larvae during early development. The mean percent time (±SE) spent displaying each of the four behavioral classifications is depicted for each day (listed at the top) and each cue (listed at the bottom). ANCOVA, a=0.05, * indicates significance.

24 1.4.3 Pseudodiploria strigosa During the two days that P. strigosa could be tested, larvae did not exhibit any significant differences in travel time (Figure 1.2C) or mean velocity (Figure 1.3C) depending on cue. Regarding the four behavior classifications, P. strigosa larvae did not show any significant variation between cues on day 3. However, on day 4, larvae spent more time in non-directed motion when presented with the reef cue (p=0.044) (Figure 1.6).

25 Pseudodiploria strigosa

Directed Movement Day 3 Day 4 30

0 70 *

60 Non-directed Movement

30 Small Circles

10 % time spent displaying each behavior each displaying spent time %

Stationary/Rotations

0 Reef Offshore Reef Offshore Figure 1.6: Variations in behavior of Pseudodiploria strigosa larvae during early development. The mean percent time (±SE) spent displaying each of the four behavioral classifications is depicted for each day (listed at the top) and each cue (listed at the bottom). ANCOVA, a=0.05, * indicates significance.

26 1.5 Discussion The successful settlement of planktonic reef larvae is determined by both water currents and animal behavior (Meadows and Campbell 1972; Raimondi and Morse 2000), but the relative role of each of these factors has been debated. While coral larval dispersal can be highly dependent on hydrodynamic forces (Wood et al. 2014), larvae can indeed alter their behavior in response to hydrostatic pressure (Stake and Sammarco 2003), light availability (Gleason et al. 2006), and waterborne chemical cues (Dixson et al. 2014) and this behavior can play a vital role in successful settlement (Raimondi and Morse 2000). During development of planktonic larvae, the relative role of behavior increases, as larvae become more functional and their preferences change (Dixson et al. 2011). The period of time between coral larvae fertilization and metamorphosis involves critical biochemical and physiological changes, including development of sensory cells and adhesion mechanisms (Chia and Bickell 1978; Gleason and Hofmann 2011). Metamorphic competence is the threshold at which environmental cues can trigger permanent attachment and metamorphosis and requires both the sensory capabilities of detecting external cues as well as the presence of appropriate chemical or physical cues (Gleason and Hofman 2011; Hadfield et al. 2011). Some coral larvae can remain planktonic for extended periods of time when suitable environmental cues are not present, but strategies for length of pelagic duration are variable between species (Bishop et al. 2006). This study investigated behavior of free-swimming coral larvae during the critical days leading up to metamorphic competence when the larvae were exposed to reef and offshore cues. As soon as A. palmata larvae were capable of swimming against the 70 mL min-1 flow in the flume, they displayed an unfailing preference for reef water over

27 offshore water throughout the remainder of their free-swimming period. Unlike reef fish (Dixson et al. 2011), no alteration in chemical choice was detected during their limited developmental time period. It is unknown if this behavior is universal among planktonic coral larvae, as A. palmata was the only species tested that was capable of swimming in the flume. A palmata larvae can reach settlement competency within 5 days but can remain planktonic up to 20 days post-fertilization (Baums 2005). With a relatively short free-swimming period, it may be advantageous for the larvae to remain invariably attracted to the reef. The majority of lecithotrophic coral larvae, which depend solely upon parental provisions during planktonic development, settle within a few days. However, some coral larvae can survive in the planktonic zone for upwards of 100 days (Graham et al. 2013). Coral species with extensive larval durations may display ontogenetic shifts in preference for reef water. The planktonic larval period of P. strigosa and O. annularis are unusual in opposite ways. P. strigosa can reach settlement competency in as little as 2.5 days post fertilization, and their larval duration usually lasts no longer than 8 days (Davies et al.

2017). In contrast, when the appropriate cue is present, Orbicella spp. usually settle within 5-14 days, however larvae of O. franksi, can survive up to 120 days without metamorphosing (Davies et al. 2017; Rippe et al. 2017). P. strigosa might be expected to behave similarly to A. palmata and remain attracted to reef water throughout their free-swimming period. It is possible that Orbicella spp. may avoid reef water during their early development, as they can remain in the pelagic zone longer (Davies et al. 2017; Rippe et al. 2017). However, it is equally as likely that extensive larval durations are caused by lack of desirable cues, so preference for reef water may occur as soon as it is detected.

28 The still-water behavioral analysis displayed variable results depending on cue, day post fertilization, and specific behavior. The clearest trend was observed in A. palmata; larvae spent more time traveling when only presented with offshore cues on day 3, but on day 5 and 6, the addition of reef water was related to an increase in movement. Ontogenetic increases in movement have been observed in other coral species, although cue was not taken into effect. For example, the percentage of slowly- moving Orbicella faveolata larvae increased through development, before dropping off again as the larvae neared settlement age (Vermeij et al. 2006). In this study, percent time spent traveling could be split into two distinct behaviors, directed and non-directed movement. Directed movement in coral larvae has been observed in other species, such as Dendrogyra cylindrus (Marharver et al. 2015), and is often related to vertical movement and positive geotaxis, or swimming towards the substrate. Non-directional movement was an all-encompassing term, which included such movements best described as spiraling, bouncing, and undulating. These same behaviors were noted in brooding coral larvae as well (Goffredo and

Zaccanti 2004). Some of the behaviors associated with non-directional movement could potentially be examples of benthic probing (Gleason et al. 2009)– especially “bouncing” behaviors, typified by larvae moving along the dish by hopping forward and touching down again. This behavior was observed across both the bottom of the petri dish and along the rim. Benthic probing in other studies is accompanied by a vertical shift in position from swimming in the water column to probing the substrate. An expected developmental shift in geotaxis was observed in the scleractinian coral, Oculina varicosa, as larvae transitioned to the substrate in search of settlement locations (Brooke and Young 2005). This study focused on the horizontal movement

29 of coral larvae and the water depth in the still-water assay was not deep enough to permit the examination of vertical movement. Although not evident in this study, other researchers have observed an increase in coral larvae velocity as development progressed and the larvae became more developed (Brooke et al. 2005; Stromberg and Larsson 2017). In this study, mean velocity per individual was analyzed, regardless of the percent time the larva spent moving. Larvae that spent more time stationary displayed disproportionately lower mean velocities, even if their maximum speed was high. Therefore, velocity was highly related to travel time and should be teased apart in future studies. Regardless, it is unknown whether or not interaction with a positive habitat cue would influence velocity of coral larvae although it is likely. Aquatic chemical cues propagate through the environment in a filamentous manner, whereby a larva passes through a series of individual streams of the cue, rather than a continuous wall of cue (Koehl et al. 2007). The larvae of the sea slug, Phestilla sibogae, ceased swimming and subsequently sank whenever the cue was encountered, and then resumed swimming in the absence of the cue (Koehl et al. 2007). This shift in velocity allowed for greater settlement success and may be a potential mechanism for coral larvae as they approach the turbulent boundary layer just above the substrate.

The still-water behavioral assay also analyzed the proportion of time the larvae spent stationary or revolving in place, and variations related to cue were detected. Two days after the initiation of swimming behaviors, larvae of A. palmata and O. annularis both experienced an increase in time spent stationary or revolving in place when exposed to the reef cue. This trend is a reversal of the previous day for O. annularis larvae, at which point they spent more time stationary in offshore water compared to

30 reef water. This definition of Stationary/Rotations aligns with what Miller and Mundy (2003) identifies as “testing” habitat, where larvae were loosely attached and spun in place. This behavior occurs before settlement, which involves both permanent attachment and subsequent metamorphosis (Miller and Mundy 2003). We did not statistically compare the variation in time spent stationary between developmental days, and no trend was clearly visible during the free-swimming period. However, on day 7 for A. palmata and day 5 for P. strigosa, larvae immediately attached themselves to the bottom upon entering the dish and did not move during the habitation period, marking the end of the free-swimming larval phase and end of the testing period. For the still-water analysis, proportion of time in each behavior and average velocity, regardless of behavior type, were compared between cues. It is possible that other parameters not addressed here may reveal variations in behavior across time. Common parameters of interest include the proportion of larvae that perform each behavior, rather than the proportion of time an individual spent preforming a specific behavior (Vermeij et al. 2006; Gleason et al. 2009). In this study, inter-individual variation was an influential factor in larval behavior post cue insertion and may have masked relevant behavioral trends. Future studies investigating chemical cue-mediated behaviors at the population level rather than the individual level, e.g. by assessing percent of stationary larvae rather than percent of time an individual larva remained stationary, may reveal temporal trends. Previous studies have assessed free-swimming coral larvae behavior when exposed to general reef cues, with variable results, which could be a factor of methodology or species-specific differences. Larvae of three Pacific acroporids

31 successfully discriminated between the chemical cues associated with a healthy, coral- dominated reef versus a degraded, algae-dominated reef (Dixson et al. 2014). In comparison, chemical cues from both a degraded and a healthy reef initiated benthic- probing and positive geotaxis in the planula of two Caribbean brooders (Gleason et al. 2009). General reef cue used in this study comes from a healthy, coral-dominated reef, although the specific chemical components of the general cue that play a role in larval behavior are unknown. Investigations into whether Caribbean larvae can differentiate between a healthy and degraded reef and what chemical components of these general reef cues initiate certain behaviors are areas of future study. In all, this study has revealed interesting temporal trends in the behavior of planktonic coral larvae when exposed to reef and offshore cues during their critical pre-settlement period. Further development of the still-water behavioral assay and associated analysis will aid in the understanding of the ecological significance of these behaviors. The still-water assay, while far removed from the turbulent, complex environment of the reef ecosystem for settling larvae, can facilitate observations of coral larvae strictly in response to a chemical cue, rather than physical movement of the water. Successful coral settlement is vital for long-term, reef revitalization efforts throughout the Caribbean. Investigations into the chemically-mediated behavior of coral larvae during habitat selection can inform research and management practices.

1.6 Acknowledgements I would like to thank D. Dixson, Z. Foltz, L. Johnston, S. Jones, V. Paul, L. Simpson, and J. Sneed for assistance with larval collection and rearing. I would also like to thank D. Dixson for collecting the fluming data. Additionally, I would like to

32 extend my gratitude to A. Barnett and E. Muñoz Ruiz for their help in analyzing the behavioral videos using Tracker software.

33 REFERENCES

Arai T, Kato M, Heyward A, Ikeda Y, Iizuka T, Maruyama T. 1993. Lipid composition of positively buoyant eggs of reef building corals. Coral Reefs. 12:71-75.

Baird AH, Guest JR, Willis BL. 2009. Systematic and biogeographical patterns in the reproductive biology of scleractinian corals. Annu Rev Ecol Evol Syst. 40:551- 571.

Bishop CD, Huggett MJ, Heyland A, Hodin J, Brandhorst BP. 2006. Interspecific variation of competence in marine invertebrates. The significance for comparative investigations into the timing of metamorphosis. Integr Comp Biol. 46:662-682.

Brooke S, Young CM. 2005. Embryogenesis and larval biology of the ahermatypic scleractinian Oculina varicosa. Mar Biol. 146:665-675.

Brown D. 2019. Tracker Video Analysis and Modeling Tool (Version 5.0.7) [Computer software]. http://physlets.org/tracker/

Bruckner AW. 2003. Proceedings of the Caribbean Acropora Workshop: potential application of the US Endangered Species Act as a conservation strategy, April 16-18, 2002, Miami, Florida.

Burke RD. 1983. The induction of metamorphosis of marine invertebrate larvae: stimulus and response. Can J Zool. 61:1701-1719.

Burke RD. 1986. Pheromones and the gregarious settlement of marine invertebrate larvae. B Mar Sci. 39:323-331.

Buschbaum C. 2001. Selective settlement of the barnacle Semibalanus balanoides (L.) facilitates its growth and reproduction on mussel beds in the Wadden Sea. Helgoland marine research, 55:128-134.

Chia FS. 1989. Differential larval settlement of benthic marine invertebrates. pp. 3-12. In Reproduction, genetics and distributions of marine organisms, J.S. Ryland and P.A. Tyler (eds.), Olsen & Olsen, Fredensborg.

34 Chia FS, Bickell LR. 1978. Mechanisms of larval attachment and the induction of settlement and metamorphosis in coelenterates: a review. pp.1-12. In: Settlement and metamorphosis of marine invertebrate larvae, F.S Chia, M.E. Rice (eds.), Elsevier-North-Holland Biomedical Press, New York.

Davies SW, Strader ME, Kool JT, Kenkel CD, Matz MV. 2017. Modeled differences of coral life-history traits influence the refugium potential of a remote Caribbean reef. Coral Reefs. 36:913-925.

Dixson DL, Munday PL, Pratchett M, Jones GP. 2011. Ontogenetic changes in responses to settlement cues by Anemonefish. Coral Reefs. 30:903-910.

Dixson DL, Abrego D, Hay ME. 2014. Chemically mediated behavior of recruiting corals and fishes: a tipping point that may limit reef recovery. Science. 345:892-897.

Eckman JE. 1983. Hydrodynamic processes affecting benthic recruitment. Limnol Oceanogr. 28:241-257.

Edmunds PJ, Elahi R. 2007. The demographics of a 15‐year decline in cover of the Caribbean reef coral Montastraea annularis. Ecol Monogr. 77:3-18.

Gerlach G, Atema J, Kingsford MJ, Black KP, Miller-Sims, V. 2007. Smelling home can prevent dispersal of reef fish larvae. Proc Natl Acad Sci USA. 104:858- 863.

Gleason DF, Hofman DK. 2011. Coral larvae: from gametes to recruits. J Exp Mar Biol Ecol. 408:42-57.

Gleason DF, Edmunds PJ, Gates RD. 2006. Ultraviolet radiation effects on the behavior and recruitment of larvae from the reef coral Porites astreoides. Mar Biol. 148:503-512.

Gleason DF, Danilowicz BS, Nolan CJ. 2009. Reef waters stimulate substratum exploration in planulae from brooding Caribbean corals. Coral Reefs. 28:549- 554.

Goffredo S, Zaccanti F. 2004. Laboratory observations of larval behavior and metamorphosis in the Mediterranean solitary coral Balanophyllia europaea (Scleractinia, Dendrophylliidae). B Mar Sci. 74:449-457.

Graham EM, Baird AH, Connolly SR, Sewell MA, Willis BL. 2013. Rapid declines in metabolism explain extended coral larval longevity. Coral Reefs. 32:539-549.

Hadfield MG. 1986. Settlement and recruitment of marine invertebrates: a perspective and some proposals. B Mar Sci. 39:418-425.

35 Hadfield MG, Paul VJ. 2001. Natural chemical cues for settlement and metamorphosis of marine invertebrate larvae. pp.431-461. In Marine Chemical Ecology, J. B. McClintock and B. J. Baker (eds.), CRC Press, Boca Raton, Florida.

Hadfield MG, Carpizo-Ituarte EJ, Del Carmen K, Nedved BT. 2011. Metamorphic competence, a major adaptive convergence in marine invertebrate larvae. Amer Zool. 41:1123-1131.

Harii S, Nadaoka K, Yamamoto M, Iwao K. 2007. Temporal changes in settlement, lipid content and lipid composition of larvae of the spawning hermatypic coral Acropora tenuis. Mar Ecol Prog Ser. 346:89-96.

Heyward AJ. 1987. Genetic systems and hereditary structures of reef corals. Doctoral dissertation. James Cook University of North Queensland, 116 pp.

Jenkins SR. 2005. Larval habitat selection, not larval supply, determines settlement patterns and adult distribution in two chthamalid barnacles. J Anim Ecol. 74:893-904.

JMP Pro, Version 13, SAS Institute Inc., Cary, NC, 1989-2007.

Jordan AC. 2018. Patterns in Caribbean Coral Spawning. Master's thesis. Nova Southeastern University.

Katz CH, Cobb JS, Spaulding M. 1994. Larval behavior, hydrodynamic transport, and potential offshore-to-inshore recruitment in the American lobster, Homarus americanus. Mar Ecol Prog Ser. 103:265-273.

Knight-Jones EW. 1953. Laboratory experiments on gregariousness during setting in Balanus balanoides and other barnacles. J Exp Biol. 30:584-598.

Knowlton N, Brainard RE, Fisher R, Moews M, Plaisance L, Caley MJ. 2010. Coral reef biodiversity. pp.65-74. In Life in the world’s oceans: diversity distribution and abundance, A. McIntyre (ed.), Oxford: Wiley-Blackwell.

Koehl MAR, Strother JA, Reidenbach MA, Koseff JR, Hadfield MG. 2007. Individual-based model of larval transport to coral reefs in turbulent, wave- driven flow: behavioral responses to dissolved settlement inducer. Mar Ecol Prog Ser. 335:1-18.

Maida M, Coll JC, Sammarco PW. 1994. Shedding new light on scleractinian coral recruitment. J Exp Mar Biol Ecol. 180:189-202.

Marhaver KL, Vermeij MJ, Medina MM. 2015. Reproductive natural history and successful juvenile propagation of the threatened Caribbean Pillar Coral Dendrogyra cylindrus. BMC Ecol. 15:1-11.

36 Meadows PS, Campbell JI. 1972. Habitat selection by aquatic invertebrates. In Advances in marine biology (Vol. 10, pp. 271-382). Academic Press.

Miller K, Mundy C. 2003. Rapid settlement in broadcast spawning corals: implications for larval dispersal. Coral Reefs. 22:99-106.

Moberg F, Folke C. 1999. Ecological goods and services of coral reef ecosystems. Ecol Econ. 29:215-233.

Mundy CN, Babcock RC. 1998. Role of light intensity and spectral quality in coral settlement: implications for depth-dependent settlement? J Exp Mar Biol Ecol. 223:235-255.

National Marine Fisheries Service. 2006. Endangered and threatened species: Final listing determinations for Elkhorn coral and Staghorn coral. Fed Regist 71:26852–26872.

National Marine Fisheries Service. 2014. Endangered and Threatened Wildlife and Plants: Adding 20 Coral Species to the List of Endangered and Threatened Wildlife. Fed Regist 79:67356-67359

Pawlik JR. 1992. Chemical ecology of the settlemenet of benthic marine invertebrates. Oceanogr Mar Biol Ann Rev. 30:273-335.

Price N. 2010. Habitat selection, facilitation, and biotic settlement cues affect distribution and performance of coral recruits in French Polynesia. Oecologia. 163:747-758.

R Core Team. 2017. R: A language and environment for statistical computing. R Foundation for Statistical Computing, Vienna, Austria. URL https://www.R- project.org/.

Raimondi PT, Morse, A.N. 2000. The consequences of complex larval behavior in a coral. Ecology. 81:3193-3211.

Rippe JP, Matz MV, Green EA, Medina M, Khawaja NZ, Pongwarin T., Pinzón C JH, Castillo KD, Davies SW. 2017. Population structure and connectivity of the mountainous star coral, Orbicella faveolata, throughout the wider Caribbean region. Ecol Evol. 7:9234-9246.

Ritson-Williams R, Paul VJ, Arnold SN, Steneck RS. 2010. Larval settlement preferences and post-settlement survival of the threatened Caribbean corals Acropora palmata and A. cervicornis. Coral Reefs. 29:71-81.

37 Ritson-Williams R, Arnold SN, Paul VJ. 2016. Patterns of larval settlement preferences and post-settlement survival for seven Caribbean corals. Mar Ecol Prog Ser. 548:127-138.

Rodríguez SR, Ojeda FP, Inestrosa NC. 1993. Settlement of benthic marine invertebrates. Mar Ecol Prog Ser. 97:193-207.

Sharp KH, Ritchie KB, Schupp PJ, Ritson-Williams R, Paul VJ. 2010. Bacterial acquisition in juveniles of several broadcast spawning coral species. PloS one. 5:e10898.

Stake JL, Sammarco PW. 2003. Effects of pressure on swimming behavior in planula larvae of the coral Porites astreoides (Cnidaria, Scleractinia). J Exp Mar Biol Ecol. 288:181-201.

Strömberg SM, Larsson AI. 2017. Larval behavior and longevity in the cold-water coral Lophelia pertusa indicate potential for long distance dispersal. Front Mar Sci 4:1-14.

Szmant AM. 1986. Reproductive ecology of Caribbean reef corals. Coral Reefs. 5:43- 53.

Tamburri MN, Finelli CM, Wethey DS, Zimmer-Faust RK. 1996. Chemical induction of larval settlement behavior in flow. Biol Bull. 191:367-73.

Tebben J, Motti CA, Siboni N, Tapiolas DM, Negri AP, Schupp PJ, Kitamura M, Hatta M, Steinberg PD, Harder T. 2015. Chemical mediation of coral larval settlement by crustose coralline algae. Sci Rep. 5:10803.

Tran C, Hadfield MG. 2013. Localization of sensory mechanisms utilized by coral planulae to detect settlement cues. Invertebr Biol. 132:195-206.

Vermeij MJA, Fogarty ND, Miller MW. 2006. Pelagic conditions affect larval behavior, survival, and settlement patterns in the Caribbean coral Montastraea faveolata. Mar Ecol Prog Ser. 310:119-128.

Vermeij MJ, Marhaver KL, Huijbers CM, Nagelkerken I, Simpson SD. 2010. Coral larvae move toward reef sounds. PloS one. 5:e10660.

Wood S, Paris CB, Ridgwell A, Hendy EJ. 2014. Modelling dispersal and connectivity of broadcast spawning corals at the global scale. Glob Ecol Biogeogr. 23:1-11.

38 Chapter 2

CORAL LARVAE BEHAVIORAL RESPONSE TO BENTHIC CUES DURING HABITAT SELECTION

2.1 Abstract Environmental chemical cues relay valuable information about the health of an ecosystem to potential larval recruits. Coral larvae recognize not only general reef cues but also those of specific benthic components of the reef ecosystem. Here, the behavioral response of Acropora palmata larvae to the chemical cues released from benthic algae and cyanobacteria was investigated using a still-water assay. Video recordings of larval behavior before and after the cue was inserted allowed for an analysis of minute behavioral changes in response to algal and cyanobacterial cues. Overall, larvae increased the time they spent traveling, regardless of the cue that was presented, but most significantly when introduced to Dictyota sp, Halimeda tuna, Galaxaura rugosa, Paragoniolithon solubile, and Hydrolithon boergesenii. Four broad behavioral classifications were identified during the assay: Stationary/Rotations, Small Circles, Non-directed Movement, and Directed Movement. Notably, the percent of time spent in directed motion or swimming in small circles significantly increased when the larvae were presented with positive cues of crustose coralline algae. These behaviors may indicate recognition of a positive settlement cue, highlighting an area for future study. Identifying positive and negative chemical cues for settling coral larvae can provide valuable insights into conservation strategies in a rapidly changing world.

39 2.2 Introduction Coral reefs are intricately complex ecosystems, replete with mobile fish and invertebrates as well as abundant benthic species of corals, algae, and sponges. All of these organisms release chemicals into their environment, either passively or actively (Hay 1996). These species-specific chemicals create an amalgam of cues that can be perceived as a general reef cue by many marine animals (Atema et al. 2002). This general reef cue reflects the species composition of the ecosystem and relays important information to potential incoming settlers regarding its quality (Dixson et al. 2014). For example, pelagic reef fish larvae can distinguish between the chemical signatures of reefs that are algal versus coral-dominated and significantly prefer healthy, coral-dominated reefs (Lecchini et al. 2013). Pelagic larvae, as well as migrating adults, can use these broad reef cues to not only orient back to a reef, but also to choose the healthiest reef in the region. The overall chemical signature of many coral reefs is changing as species distributions and abundances shift in response to environmental stressors (Nyström et al. 2000; Hughes et al. 2003). Overfishing, eutrophication, bleaching, and structural damage can result in phase shifts characterized by an alteration in the dominant benthic species on a reef (Hughes 1994; Bruno et al. 2009). For example, in the 1980s, the Caribbean experienced massive mortality rates of the long-spined sea urchin, Diadema antillarum, a grazer vital in controlling marine macroalgae (Lessios 1988). The functional extinction of this species, along with other local stressors, led to a rapid shift from coral-dominated reef systems to algal-dominated systems (Liddell and Ohlhorst 1986). Progress towards recovery has occurred, but it has been extremely slow (Lessios 2016).

40 As Caribbean reefs continue to experience phase shifts towards algal- domination, with limited capacity for recovery (Mumby et al. 2007), research has focused on the impact of benthic algae on reef-building corals throughout their life history, from fertilization to sexual maturity (Birrell et al. 2008a). Within the reef matrix, upright macroalgae of many different taxonomic and functional groups directly compete with settled corals for the limited available substrate on which to grow (McCook et al. 2001). Generally, both corals and benthic algae require solid substrate to anchor themselves as well as unobstructed sunlight for photosynthesis. Mechanisms for competition between algae and corals include overgrowth, shading, abrasion, epithelial sloughing, recruitment barriers, and chemical defense (McCook et al. 2001). In this race for both settlement sites and light, many benthic species produce defensive compounds that are repulsive or toxic to potential consumers (Hay and Fenical 1988). Further, chemicals are also used for direct competition with neighbors through allelopathic warfare (Littler and Littler 1997; Gross 2003; Rasher and Hay 2010, 2014; Rasher et al. 2011).

Settlement in an area replete with upright benthic algae can be detrimental to corals of all sizes, therefore pre-settlement larvae should aim to avoid these areas. Mobile coral larvae are capable of recognizing environmental cues and can modify their behavior in response to these cues (Dixson et al. 2014). Detection of external cues is a requirement of metamorphic competency and is thus a critical aspect of early development (Gleason and Hofman 2011; Hadfield et al. 2011). Research has shown that the presence of cues from a host of upright benthic macroalgae and cyanobacteria can deter settlement in coral larvae (Kuffner et al. 2006; Birrell et al. 2008a), yet research into specific behavioral responses before settlement is limited.

41 While the presence of some macroalgal cues can deter settlement, other cues can induce settlement. Crustose coralline algae (CCA) are a group of calcareous red algae that act as cement in a reef ecosystem (Steneck 1986; Adey 1998) and contain chemical cues, potentially in their associated biofilm, that are attractive to coral larvae (Morse et al. 1988; Heyward and Negri 1999; Hadfield and Paul 2001; Negri et al. 2001; Harrington et al. 2004; Golbuu and Richmond 2007; Kitamura et al. 2007, 2009; Sneed et al. 2014; Tebben et al. 2015). Larval preference for CCA is species specific as CCA growth forms can vary, with some species providing solid substrate ideal for settling larvae and other species being unsuitable for permanent settlement due to epithelial sloughing (Harrington et al. 2004). It would be beneficial for free-swimming coral larvae within the reef matrix to distinguish between chemicals produced by different reef algae, as an inappropriate settlement choice can lead to decreased fitness or death. The species-specific chemical preferences of free-swimming coral larvae has been investigated in the Indo-Pacific using a two-channel choice flume (Dixson et al.

2014). Coral larvae are not only capable of using general reef cues to distinguish between healthy and degraded reefs, but they also respond to species-specific benthic cues of corals and algae. Coral larvae are chemically attracted to CCA and adult corals but are repulsed by several species of macroalgae (Dixson et al. 2014). Research into pre-settlement behavioral response of coral larvae to algal cues is ongoing in the Caribbean. The current study attempts to determine the minute changes in behavior of Caribbean coral larvae after exposure to common algal and cyanobacteria cues. I developed a novel still-water behavioral assay to investigate the response of Acropora palmata larvae to common Caribbean macroalgae and

42 cyanobacteria of various taxonomic and functional groups. As settlement is permanent after metamorphosis and the consequences of inappropriate settlement sites are high, I hypothesized that the introduction of benthic cues, either positive or negative, would incite a behavioral reaction in free-swimming coral larvae. Specifically, due to the detrimental effects of upright macroalgae on corals at all life stages and the benefits of CCA for inducing metamorphosis, I hypothesized that coral larvae would display distinctive behaviors when presented with traditionally negative versus positive cues.

2.3 Methods

2.3.1 Larval collection and rearing

This study was conducted at the Carrie Bow Cay field station, off the coast of Belize (16°48'9.26"N, 88° 4'54.87"W). Acropora palmata is a critically-endangered, yet vital reef-building coral throughout the Caribbean (see Chapter 1 methods for species information). Acropora palmata coral larvae were collected from the reefs surrounding Carrie Bow Cay, Belize on 5-6 August 2015 and 23-24 August 2016 according to methods described in Chapter 1. Fertilized eggs were reared in 4 L larval containers with continual seawater flow (Ritson-Williams et al. 2010). Throughout development, deceased larvae were removed from the container using pipettes. Larvae were reared for at least 4 days before initiating behavioral assays. Assays were run on larvae 4-6 days post spawn, after which the larvae ceased swimming in behavior trials and remained attached to the substrate.

2.3.2 Cue generation Common Caribbean algal and cyanobacterial species of various taxonomic and functional groups (Table 2.1) were used as environmental cues in the still-water assays

43 and were collected from the barrier reef around Carrie Bow Cay in waters ranging from 1-15 m. Macroalgae (Galaxaura rugosa, Halimeda tuna, H. opuntia, Stypopodium zonale, Dictyota sp., and Lobophora sp.) and a cyanobacterium (Hormothamnion enteromorphoides) were semi-dried in a salad spinner for 25 revolutions to reduce excess water before being weighed to 20 g (Dixson 2011). Crustose coralline algae (Paragoniolithon solubile and Hydrolithon boergesenii) were collected from reef rubble (1 m depth) using rock hammers and trimmed with wire clippers to eliminate additional cues before being weighed to 50 g. After being weighed to the right amount, cues were individually soaked in 10 L seawater for 1 hour (Dixson 2011) to generate the cues for the experiments.

44 Table 2.1: Functional and taxonomic groups of benthic algae and cyanobacteria used in this study (from 1Steneck and Dethier 1994, 2Littler et al. 1983, 3Diaz-Pulido 2010).

Species Functional group Taxonomic group

1Corticated foliose, Stypopodium zonale 2Thick leathery group 3Upright, fleshy Phaeophyceae (Brown algae) Lobophora sp. 1Corticated foliose, 2Sheet group Dictyota sp. 3Upright, fleshy

Halimeda tuna 1Articulated calcareous, Chlorophyta (Green algae) Halimeda opuntia 2Jointed calcareous group 3Upright, calcareous Galaxaura rugosa

Hydrolithon boergesenii Rhodophyta (Red algae) 1,2,3Crustose calcareous Paragoniolithon solubile Hormothamnion 1Microalgae Cyanophyta (Cyanobacteria) enteromorphoides

2.3.3 Still-water behavioral assay

A still-water assay was used to assess minute changes in coral larvae behavior when exposed to algal cues. As described in more detail in chapter 1, a larva was pipetted into a petri dish filled with 30mL filtered seawater. After a 1 min habituation period, video recording above the dish commenced. After 2 min, the larva was pipetted back to the center of the dish and 2 mL of cue water and 2 mL of control water were carefully inserted into the far sides of the dish. Video recording continued for 2 min before the conclusion of the trial.

45 2.3.4 Video analysis Average velocity was analyzed using Tracker 5.0 (Brown 2019), as described in chapter 1. Additionally, videos were analyzed manually to characterize behavior type. Four general behaviors were identified: (1) Stationary Rotations, in which the larva either remained motionless or rotated in place, (2) Small Circles, in which the larva swam in circles <1cm, yet did not experience any overall change in position within the dish, (3) Directed Movement, in which the larva swam in one direction with limited variations in speed; and (4) Non-directed Movement, in which the larva traveled in an unpredictable fashion involving spiraling, drifting, bouncing, and serpentine motions. For both Tracker and manual analyses, the first 10 seconds of footage after the cue was introduced was removed to account for any water movement due to the induction of the cue rather than larval self-propulsion.

2.3.5 Statistics

This study investigated behavioral changes of coral larvae after a benthic cue was presented; therefore, analyses focused on the differences in pre-cue and post cue data. Mean velocity and percent time spent displaying each behavior were analyzed using the Wilcoxon signed rank test (JMP Pro 13), a non-parametric alternative to the paired t-test as the data was not normally distributed despite transformations.

2.4 Results

2.4.1 Brown algae Three species of brown algae were tested in the still-water assays, Stypopodium zonale, Dictyota sp., and Lobophora sp. Neither the introduction of S. zonale nor Lobophora sp. chemical cues resulted in significant variations in larval

46 velocity or percentage of time spent in any particular behavior. Only the introduction of chemical cues from Dictyota sp. resulted in a significant increase in the time the larvae spent traveling (p=0.025, Figure 2.1) as well as average larval velocity (p<0.001, Figure 2.2). Additionally, a significant decrease in stationary/rotation behavior (p=0.005, Figure 2.3A) was paired with an increase in non-directed movement (p=0.023, Figure 2.3C) after Dictyota sp. cue was presented to the coral larvae.

100 Post-cue 90 Pre-cue 80

70 *

60 * * * 50 *

30 % time spent traveling spent time %

0 . . sp sp

Dictyota Lobophora Halimeda tuna Stypopodium zonale Halimeda opuntiaGalaxaura rugosa HydrolithonParagoniolithon boergesenii solubile

Hormothamnion enteromorphoides

47 0.25 Post-cue Pre-cue 0.2

0.15 * * 0.1 *

Mean velocity (cm/s) velocity Mean *

0.05

0 sp. sp.

Dictyota Lobophora Halimeda tuna Stypopodium zonale Halimeda opuntiaGalaxaura rugosa Hydrolithon boergeseniiParagoniolithon solubile

Hormothamnion enteromorphoides

48 A. Stationary/Rotations B. Small Circles * Post-cue 80 * * 80 * Pre-cue

60 * 60

40 40 % time time % * 20 20

0 0 C. Non-Directed Movement D. Directed Movement 80 80

60 60 * * 40 * 40 % time time % * 20 20 *

0 0 . sp sp.

Dictyota sp. Dictyota Lobophora sp. Halimeda tuna Lobophora Halimeda tuna HalimedaGalaxaura opuntia rugosa HalimedaGalaxaura opuntia rugosa Stypopodium zonale Stypopodium zonale HydrolithonParagoniolithon boergesenii solubile HydrolithonParagoniolithon boergesenii solubile

Hormothamnion enteromorphoides Hormothamnion enteromorphoides Figure 2.3: Comparison of pre-cue and post-cue mean percent time (±SE) spent in each of the four behavioral classifications, (A) Stationary/Rotations, (B) Small Circles, (C) Non-Directed Movement, and (D) Directed Movement. Lighter colors refer to pre-cue data and darker colors refer to post-cue data. Brown bars=brown algae, green bars=green algae, red bars=upright red algae, blue bars= cyanobacteria, purple bars=CCA. Wilcoxon signed rank test, a=0.05, * indicates significance.

2.4.2 Green algae

Analysis of the still-water behavioral videos revealed no change in coral larvae behavior when presented with chemical cues of H. opuntia. However, when the larvae were presented with H. tuna cues, they exhibited an increase in percent time traveling (p=0.008, Figure 2.1) and an associated increase in velocity (p=0.035, Figure 2.2). Specifically, an increase in the percent time in non-directed motion (p=0.004, Figure

49 2.3C) was reflected in a decrease in the percent time stationary or revolving in place (p=0.005, Figure 2.3A) after the cue was presented.

2.4.3 Red algae Galaxaura rugosa was the only species of upright red algae used in the behavioral assays. A. palmata larvae significantly increased their percent time spent traveling when presented with the chemical cues of G. rugosa (p=0.018, Figure 2.1), but an associated increase in average velocity was not detected (Figure 2.2). Larvae significantly increased the percent time spent in non-directed motion (p=0.005, Figure 2.3C) and reduced the percent time spent stationary or revolving in place (p=0.026, Figure 2.3A) after exposure to G. rugosa cues.

2.4.4 Cyanobacteria

Hormothamnion enteromorphoides acted as a representative cyanobacterium during the still-water trials. No significant changes in mean velocity or behavior were detected when A. palmata larvae were presented with cues from H. enteromorphoides.

2.4.5 Crustose coralline algae (CCA) The addition of chemical cues from the two CCA species, P. solubile and H. boergesenii, into the petri dishes of the still-water assays resulted in a significant increase in coral larvae travel time (p<0.001, Figure 2.1) as well as mean velocity (H. boergesenii: p<0.001, P. solubile: p=0.048, Figure 2.2). Larvae consistently displayed a decrease in time spent stationary or revolving in place after the cues were inserted (H. boergesenii: p<0.001, P. solubile: p=0.002, Figure 2.3A). The addition of cues from P. solubile was the only alga to cause an increase in amount of time spent swimming in small circles (p=0.012, Figure 2.3B). When waterborne cues of H.

50 boergesenii were presented, coral larvae displayed a significant increase in both directed and non-directed movement (p=0.012, Figure 2.4D; p=0.007, Figure 2.4C, respectively)

2.5 Discussion Upright macroalgae and cyanobacteria can directly compete with coral larvae for space on the substrata to settle, therefore avoidance of chemical cues associated with these species would be beneficial for coral fitness. Additionally, benthic algae and cyanobacteria from a variety of taxonomic and functional groups produce compounds that can be detrimental for survival and recruitment of invertebrate larvae (Walters et al. 1996), including coral larvae (Diaz-Pulido et al. 2010). Numerous studies have detected a reduction in survival and/or settlement of coral larvae after exposure to chemical cues of brown algae (Morse 1996; Baird and Morse 2004; Kuffner 2006; Birrell 2008b; Vermeij et al. 2009; Diaz-Pulido et al. 2010; Paul et al. 2011; Fong et al. 2019), green algae (Birrell et al. 2008b; Vermeij et al. 2009; Diaz- Pulido et al. 2010; Lee et al. 2012), upright red algae (Vermeij et al. 2009; Diaz-Pulido et al. 2010), and cyanobacteria (Kuffner and Paul 2004; Kuffner 2006). However, interactions are not consistent and various species of coral larvae respond positively or indifferently to cues of some upright benthic algae and cyanobacteria (Maypa and Raymundo 2004; Nugues and Szmant 2006; Birrell 2008b). Encrusting red algae, or the associated biofilm, is a known attractant to coral larvae and can induce settlement and metamorphosis (Morse et al. 1988; Heyward and Negri 1999; Hadfield and Paul 2001; Negri et al. 2001; Harrington et al. 2004; Webster et al. 2004; Golbuu and Richmond 2007; Kitamura et al. 2007, 2009; Ritson-Williams et al. 2010; Sneed et al.

51 2014; Tebben et al. 2015); however, compounds have been derived from some CCA that are toxic to coral larvae (Kitamura et al. 2005). Most of the above studies assessed settlement success, but pre-settlement coral larvae are also capable of altering their swimming direction to reflect their preference (Dixson et al. 2014). In the Indo-Pacific, Acropora tenuis larvae swam towards cues associated with CCA and away from chemical cues of upright brown and red algae (Dixson et al. 2014). Movement towards or away from a cue may not be the only behavioral response initiated by a coral larva after exposure to a repulsive or attractive cue. Coral larvae have been observed performing a diversity of behaviors including, bouncing, spiraling, rotating in place, and swimming in small circles (Miller and Mundy 2003; Goffredo and Zaccanti 2004; Gleason et al. 2009; Marharver et al. 2015). This study assessed behavioral changes of coral larvae when presented with common Caribbean algal cues, and these deviations were variable depending on the cue, even within algal groups. Overall, the introduction of algal cues resulted in decreased stationary/rotations and increased non-directional movement. This trend, most apparent when the larvae were exposed to Dictyota sp., H. tuna, G. rugosa, and the CCA species, P. solubile and H. boergesenii, and was unrelated to algal taxonomic or functional relatedness.

While CCA typically induce settlement, upright brown, red, and green algae typically deter settlement (Birrell et al. 2008a). An increase in movement regardless of cue may indicate that the sensory capabilities of coral larvae are developed enough to perceive environmental chemicals. Additionally, coral larvae may utilize the same behavioral mechanisms to both avoid or swim towards a cue.

52 Notably, the increase in the proportion of time spent in small circles or directed movement is unique to the introduction of potentially positive cues. Coral larvae spent more time performing small circles after the CCA, P. solubile, was introduced. Swimming in uniform small circles, without changing overall position within the dish was a very distinct, but relatively rare behavior. Bryozoan larvae display this behavior when an appropriate settlement site has been located (Ryland 1974; Chia et al. 1983), suggesting that swimming in tight circles might be indicative of a positive cue for coral larvae settlement as well. Additionally, zooplankton utilize a similar pattern of cyclical loops when they encounter a high-density food patch (Williamson 1981; Hamner et al. 1983; Buskey 1984). Swimming in small circles allows zooplankton such as copepods to remain longer within the area of abundant food resources. Coral larvae may engage in cyclical swimming in an attempt to remain near the source of a positive cue so that settlement can occur. Swimming in small circles was also distinct but rare throughout the development of A. palmata, Orbicella annularis, and Pseudodiploria strigosa larvae (see Chapter 1). Further investigations into the nature and prevalence of this behavior would likely prove insightful in the identification of positive settlement cues. A. palmata larvae increased the amount of time traveling in a directed motion only when the CCA, H. boergesenii, was detected, although the directionality of that movement was not assessed. H. boergesenii has been shown to be a settlement inducer for several species of coral larvae (Ritson-Williams et al. 2010, 2016) and in a paired choice settlement assay, A. palmata preferred to settle onto H. boergesenii over P. solubile (Ritson-Williams et al. 2010). An increase in directed motion after the addition of H. boergesenii cues indicates that directed motion might be indicative of a

53 positive benthic settlement cue. Directed motion was observed in larvae of the Pillar coral, Dendrogyra cylindrus, while the larvae were positively geotactic and swimming near the bottom of the container (Marharver et al. 2015). The shallow nature of our testing arena revealed directed motion along a horizontal plane, but vertical directed motion may be evident in deeper containers, especially as larvae reach settlement age. Of the brown algae, only Dictyota sp. cues triggered significant variations in larval behavior in the still-water analyses, despite all three Phaeophyceae species containing significant quantities of secondary metabolites, some known to inhibit coral settlement (Gerwick et al. 1985; Kuffner et al. 2006; Diaz-Pulido et al. 2010; Paul et al. 2011; Fong et al. 2019). Similarly, introduction of the chemical cues of the cyanobacterium, H. enteromorphoides, did not incite a significant reaction in coral larvae during the behavioral assays, despite similar cyanobacteria species deterring settlement of other scleractinian coral species (Kuffner and Paul 2004; Kuffner 2006). These upright macroalgae and cyanobacteria are abundant on degraded reefs, leading us to incorrectly hypothesize that cues associated with these species would cause a reaction in coral larval behavior in the still-water assay. Perceived indifference to waterborne cues of Lobophora sp., S. zonale, and H. enteromorphoides in the still- water trials could potentially be due to high inter-individual variation or lack of cue detection but could also indicate that these benthic species do not emit compounds that are relevant to coral larvae. Results of this study corroborate previous studies suggesting that response to benthic cues depends on the focal larval species and remains highly variable regardless of algal taxonomic or functional relatedness. For example, H. opuntia inhibits settlement of the brooder, Pocillopora damicornis (Lee et al. 2012), yet larvae

54 of Favia fragrum surprisingly preferred to settle directly on H. opuntia individuals (Nugues and Szmant 2006). In this study, A. palmata larvae were indifferent to H. opuntia cues, but increased movement and velocity after exposure to chemical cues of the congeneric, H. tuna. Similarly, A. palmata larvae were indifferent to cues of H. enteromorphoides, despite this species producing secondary metabolites deterrent to herbivores (Pennings et al. 1997). However, the presence of this cyanobacterium was correlated to an increase in successful recruitment of coral larvae in Hawaii (Cover 2011). This correlation, however, may be an indirect result of a reduction in grazers that scrape off coral larvae due to the presence of the cyanobacterium. Therefore, even though the specific chemical cues of these species deter grazers, they might reflect potentially beneficial macrohabitat, resulting in conflicting responses. Definitive avoidance of macroalgal chemical cues may offer an evolutionary advantage to coral larvae for two reasons. First, secondary metabolites produced by benthic algae may directly inhibit coral growth (Tanner 1995; Lirman 2001; Rasher and Hay 2010, 2014; Rasher et al. 2011). Second, even if some algal cues are not acutely toxic, they may still relay important information about potential settlement locations at both the macro ecosystem scale and the microhabitat scale. Within the reef matrix, it would be disadvantageous for a coral larva to settle adjacent to a fast- growing, chemically-rich alga with the potential to shade sunlight. Additionally, as Caribbean reefs rapidly shift from coral-dominated to algae-dominated, an abundance of algal cues may warn free-swimming coral larvae of a degraded reef (Lecchini et al. 2013). In conclusion, observing and quantifying the behavioral patterns of coral larvae is an important step in coral larval ecology and requires the use of diverse and novel

55 assays that can be adapted based on coral species, relevant environmental cues, and behavioral traits of interest. As behavioral patterns can influence recruitment success (Raimondi and Morse 2000), determining how coral larvae will respond to environmental cues can be vital for conservation efforts. Identification of positive and negative chemical cues that impact recruitment and settlement of coral larvae can inform reef management. For example, herbivores that consume repulsive algae can be granted high protection status (Bellwood et al. 2004), and settlement-inducing chemicals can be incorporated into artificial reefs. An interdisciplinary approach to reef management, incorporating aspects of animal behavior and environmental cues, can allow for more efficient, ecosystem-level conservation.

2.6 Acknowledgements I would like to thank D. Dixson, Z. Foltz, L. Johnston, S. Jones, V. Paul, L. Simpson, and J. Sneed for assistance with larval collection and rearing. I would additionally like to thank V. Paul and J. Sneed for collection and identification of algae and cyanobacteria in the field. Finally, I would express my gratitude towards A. Barnett, M. Brennan, M. Kitchen, and C. McCrone for their help in analyzing the behavioral videos using Tracker software.

56 REFERENCES

Adey WH. 1998. Coral reefs: algal structured and mediated ecosystems in shallow, turbulent, alkaline waters. J Phycol. 34:393-406.

Atema J, Kingsford MJ, Gerlach G. 2002. Larval reef fish could use odour for detection, retention and orientation to reefs. Mar Ecol Prog Ser. 4:151-160.

Baird AH, Morse ANC. 2004. Induction of metamorphosis in larvae of the brooding corals Acropora palifera and Stylophora pistillata. Mar Freshwater Res. 55:469–472.

Bellwood DR, Hughes TP, Folke C, Nyström M. 2004. Confronting the coral reef crisis. Nature. 429:827-833.

Birrell CL, McCook LJ, Willis BL, Diaz-Pulido GA. 2008a. Effects of benthic algae on the replenishment of corals and the implications for the resilience of coral reefs. Oceanogr Mar Biol Annu Rev. 46:25-63.

Birrell CL, McCook LJ, Willis BL, Harrington L. 2008b. Chemical effects of macroalgae on larval settlement of the broadcast spawning coral Acropora millepora. Mar Ecol Prog Ser. 362:129-137.

Brown D. 2019. Tracker Video Analysis and Modeling Tool (Version 5.0.7) [Computer software]. http://physlets.org/tracker/

Bruno JF, Sweatman H, Precht WF, Selig ER, Schutte VG. 2009. Assessing evidence of phase shifts from coral to macroalgal dominance on coral reefs. Ecology. 90:1478-1484.

Buskey EJ. 1984. Swimming pattern as an indicator of the roles of copepod sensory systems in the recognition of food. Marine Biology. 79:165-175.

Chia FS, Buckland-Nicks J, Young CM. 1984. Locomotion of marine invertebrate larvae: a review. Can J Zool. 62:1205-22.

Cover WA. Species interactions affecting corals and recruitment on a protected, high- latitude reef: herbivory, predation, and competition by fishes, urchins, macroalgae and cyanobacteria. University of California, Santa Cruz; 2011 Jun.

57 Diaz-Pulido G, Harii S, McCook LJ, Hoegh-Guldberg O. 2010. The impact of benthic algae on the settlement of a reef-building coral. Coral Reefs. 29:203-208.

Dixson DL. 2011. Smelling home: the use of olfactory cues for settlement site selection by coral reef fish larvae. Doctoral dissertation. James Cook University.

Dixson DL, Abrego D, Hay ME. 2014. Chemically mediated behavior of recruiting corals and fishes: a tipping point that may limit reef recovery. Science. 345:892-897.

Fong J, Lim ZW, Bauman AG, Valiyaveettil S, Liao LM, Yip ZT, Todd PA. 2019. Allelopathic effects of macroalgae on Pocillopora acuta coral larvae. Mar Environ Res (In Press).

Gerlach G, Atema J, Kingsford MJ, Black KP, Miller-Sims, V. 2007. Smelling home can prevent dispersal of reef fish larvae. Proc Natl Acad Sci USA. 104:858- 863.

Gleason DF, Hofman DK. 2011. Coral larvae: from gametes to recruits. J Exp Mar Biol Ecol. 408:42-57.

Gerwick WH, Fenical W, Norris JN. 1985. Chemical variation in the tropical seaweed Stypopodium zonale (Dictyotaceae). Phytochemistry. 24:1279-83.

Gleason DF, Danilowicz BS, Nolan CJ. 2009. Reef waters stimulate substratum exploration in planulae from brooding Caribbean corals. Coral Reefs. 28:549- 554.

Golbuu Y, Richmond RH. 2007. Substratum preferences in planula larvae of two species of scleractinian corals, Goniastrea retiformis and Stylaraea punctata. Mar Biol. 152:639-644.

Gross EM. 2003. Allelopathy of Aquatic Autotrophs. Crit Rev Plant Sci. 22:313-339.

Hadfield MG, Paul VJ. 2001. Natural chemical cues for settlement and metamorphosis of marine invertebrate larvae. pp.431-461. In Marine Chemical Ecology, J. B. McClintock and B. J. Baker (eds.), CRC Press, Boca Raton, Florida.

Hadfield MG, Carpizo-Ituarte EJ, Del Carmen K, Nedved BT. 2011. Metamorphic competence, a major adaptive convergence in marine invertebrate larvae. Amer Zool. 41:1123-1131.

58 Hamner WM, Hamner PP, Strand SW, Gilmer RW. 1983. Behavior of Antarctic krill, Euphausia superba: chemoreception, feeding, schooling, and molting. Science. 220:433-435.

Harrington L, Fabricius K, De'Ath G, Negri A. 2004. Recognition and selection of settlement substrata determine post‐settlement survival in corals. Ecology. 85:3428-37.

Hay ME. 1996. Marine chemical ecology: what's known and what's next? J Exp Mar Biol Ecol. 200:103-134.

Hay ME, Fenical W. 1988. Marine plant-herbivore interactions: the ecology of chemical defense. Annu Rev Ecol Evol Syst. 19:111-145.

Heyward AJ, Negri AP. 1999. Natural inducers for coral larval metamorphosis. Coral Reefs. 18:273-279.

Hughes TP. 1994. Catastrophes, phase shifts, and large-scale degradation of a Caribbean coral reef. Science. 265:1547-1551.

Hughes TP, Baird AH, Bellwood DR, Card M, Connolly SR, Folke C, Grosberg R, Hoegh-Guldberg O, Jackson JB, Kleypas J, Lough JM. 2003. Climate change, human impacts, and the resilience of coral reefs. Science. 301:929-933.

JMP Pro, Version 13, SAS Institute Inc., Cary, NC, 1989-2007.

Kitamura M, Koyama T, Nakano Y, Uemura D. 2005. Corallinafuran and corallinaether, novel toxic compounds from crustose coralline red algae. Chem Lett. 34:1272–1273.

Kitamura M, Koyama T, Nakano Y, Uemura D. 2007. Characterization of a natural inducer of coral larval metamorphosis. J Exp Mar Biol Ecol. 340:96-102.

Kitamura M, Schupp PJ, Nakano Y, Uemura D. 2009. Luminaolide a novel metamorphosis-enhancing macrodiolide for scleractinian coral larvae from crustose coralline algae. Tetrahedron Lett. 50:6606-6609.

Kuffner IB, Paul VJ. 2004. Effects of the benthic cyanobacterium Lyngbya majuscula on larval recruitment of the reef corals Acropora surculosa and Pocillopora damicornis. Coral Reefs. 23:455-458.

Kuffner IB, Walters LJ, Becerro MA, Paul VJ, Ritson-Williams R, Beach KS. 2006. Inhibition of coral recruitment by macroalgae and cyanobacteria. Mar Ecol Prog Ser. 323:107-117.

59 Liddell WD, Ohlhorst SL. 1986. Changes in benthic community composition following the mass mortality of Diadema at Jamaica. J Exp Mar Biol Ecol. 95:271-278.

Littler MM, Littler DS, Taylor PR. 1983. Evolutionary strategies in a tropical barrier reef system: Functional-form groups of marine macroalgae 1. J Phycol. 19:229-237.

Lecchini D, Waqalevu VP, Parmentier E, Radford CA, Banaigs B. 2013. Fish larvae prefer coral over algal water cues: implications of coral reef degradation. Mar Ecol Prog Ser. 475:303-307.

Lee, C.S., Walford, J. and Goh, B., 2012. The effect of benthic macroalgae on coral settlement.

Lessios HA. 1988. Mass mortality of Diadema antillarum in the Caribbean: what have we learned? Annu Rev Ecol Syst. 19:371–393.

Lessios HA. 2016. The great Diadema antillarum die-off: 30 years later. Ann Rev Mar Sci. 8:267-283.

Lirman D. 2001. Competition between macroalgae and corals: effects of herbivore exclusion and increased algal biomass on coral survivorship and growth. Coral reefs. 19:392-399.

Littler MM, Littler DS. 1997. Epizoic red alga allelopathic (?) to a Caribbean coral. Coral Reefs. 16:168.

Marhaver KL, Vermeij MJ, Medina MM. 2015. Reproductive natural history and successful juvenile propagation of the threatened Caribbean Pillar Coral Dendrogyra cylindrus. BMC Ecol. 15:1-11.

Miller K, Mundy C. 2003. Rapid settlement in broadcast spawning corals: implications for larval dispersal. Coral Reefs. 22:99-106.

McCook L, Jompa J, Diaz-Pulido G. 2001. Competition between corals and algae on coral reefs: a review of evidence and mechanisms. Coral reefs. 19:400-17.

Morse DE, Hooker N, Morse AN, Jensen RA. 1988. Control of larval metamorphosis and recruitment in sympatric agariciid corals. J Exp Mar Biol Ecol. 116:193- 217.

Morse AN, Iwao K, Baba M, Shimoike K, Hayashibara T, Omori MA. 1996. An ancient chemosensory mechanism brings new life to coral reefs. Biol Bull. 191:149-54.

60 Mumby PJ, Hastings A, Edwards HJ. 2007. Thresholds and the resilience of Caribbean coral reefs. Nature. 450:98-101.

Negri AP, Webster NS, Hill RT, Heyward AJ. 2001. Metamorphosis of broadcast spawning corals in response to bacteria isolated from crustose algae. Mar Ecol Prog Ser. 223:121-131.

Nugues MM, Szmant AM. 2006. Coral settlement onto Halimeda opuntia: a fatal attraction to an ephemeral substrate? Coral Reefs. 25:585-591.

Nyström M, Folke C, Moberg F. 2000. Coral reef disturbance and resilience in a human-dominated environment. Trends Ecol Evol. 15:413-7.

Paul VJ, Fenical W. 1983. Isolation of halimedatrial: chemical defense adaptation in the calcareous reef-building alga Halimeda. Science. 221:747-749.

Paul VJ, Van Alstyne KL. 1988. Chemical defense and chemical variation in some tropical Pacific species of Halimeda (Halimedaceae; Chlorophyta). Coral Reefs. 6:263-9.

Paul VJ, Kuffner IB, Walters LJ, Ritson-Williams R, Beach KS, Becerro MA. 2011. Chemically mediated interactions between macroalgae Dictyota spp. and multiple life-history stages of the coral Porites astreoides. Mar Ecol Prog Ser. 426:161-170.

Pawlik JR. 1992. Chemical ecology of the settlement of benthic marine invertebrates. Oceanogr Mar Biol Annu Rev. 30:273-335.

Pennings SC, Pablo SR, Paul VJ. 1997. Chemical defenses of the tropical, benthic marine cyanobacterium Hormothamnion enteromorphoides: Diverse consumers and synergisms. Limnol Oceanogr. 42:911–917.

Rasher DB, Hay ME. 2010. Seaweed allelopathy degrades the resilience and function of coral reefs. Commun Integr Biol. 3:564-566.

Rasher DB, Hay ME. 2014. Competition induces allelopathy but suppresses growth and anti-herbivore defense in a chemically rich seaweed. Proc R Soc Lond B Biol Sci. 281:20132615.

Rasher DB, Stout EP, Engel S, Kubanek J, Hay ME. 2011. Macroalgal terpenes function as allelopathic agents against reef corals. Proc Natl Acad Sci USA. 108:17726-17731.

Raimondi PT, Morse, A.N. 2000. The consequences of complex larval behavior in a coral. Ecology. 81:3193-3211.

61 Ritson-Williams R, Paul VJ, Arnold SN, Steneck RS. 2010. Larval settlement preferences and post-settlement survival of the threatened Caribbean corals Acropora palmata and A. cervicornis. Coral Reefs. 29:71-81.

Ritson-Williams R, Arnold SN, Paul VJ. 2016. Patterns of larval settlement preferences and post settlement survival for seven Caribbean corals. Mar Ecol Prog Ser. 548:127-138.

Ryland JS. 1974. Behavior, settlement and metamorphosis of bryozoan larvae: a review. Thalassia Jugosl. 10:239-262.

Steneck RS. 1986. The ecology of coralline algal crusts: convergent patterns and adaptive strategies. Annu Rev Ecol Evol Syst. 17:273-303.

Steneck RS, Dethier MN. 1994. A functional group approach to the structure of algal- dominated communities. Oikos. 69:476-498.

Tanner JE. 1995. Competition between scleractinian corals and macroalgae: an experimental investigation of coral growth, survival and reproduction. J Exp Mar Biol Ecol. 190:151-68.

Vermeij MJ, Smith JE, Smith CM, Thurber RV, Sandin SA. 2009. Survival and settlement success of coral planulae: independent and synergistic effects of macroalgae and microbes. Oecologia. 159:325-336.

Walters LJ, Hadfield MG, Smith CM. 1996. Waterborne chemical compounds in tropical macroalgae: positive and negative cues for larval settlement. Mar Biol. 126:383-93.

Williamson CE. 1981. Foraging behavior of a freshwater copepod: frequency changes in looping behavior at high and low prey densities. Oecologia. 50:332-336.

62 Chapter 3

MULTIPLE ENVIRONMENTAL CUES IMPACT HABITAT CHOICE DURING NOCTURNAL HOMING OF SPECIALIZED REEF SHRIMP

This chapter has been published in the journal Behavioral Ecology. Co-Authors: Danielle L. Dixson1 Addresses: 1School of Marine Science and Policy, University of Delaware, 700 Pilottown Road, Lewes, DE, 19958, USA

3.1 Abstract Habitat selection is a critical process for animals throughout their life, and adult organisms that travel to forage or mate must re-select habitat frequently. On coral reefs, competition for space has led to a high proportion of habitat specialists. Habitat selection is especially vital for organisms that require specialized habitat; however, research has primarily focused on the initial habitat choice made during the larval/juvenile stage. Here, we analyze habitat selection in the adult sponge-dwelling reef shrimp, Lysmata pederseni. Using a mark-and-recapture technique, belt transects, patch reefs, and cue isolation experiments, this study reveals that adult L. pederseni diurnally re-select habitat and a natural preference exists for specific sponge species and shapes. This natural preference is a function of chemical and morphological cues as well as sponge distribution. As habitat specialists can drive biodiversity, understanding the mechanisms behind habitat selection can inform research and management practices.

63 3.2 Introduction Animals typically occupy distinct habitats ranging in specificity from the entire ecosystem to specialized single-host microhabitats. Habitat selection has been researched in a variety of taxa in an attempt to understand the balance between environmental variables (Cowen and Sponaugle 2009) and animal behavior (Meadows and Campbell 1972). While some organisms choose appropriate habitat once during their life history – typically at the juvenile or larval stage – others must re-select habitat frequently if they migrate, disperse, or temporally home. Roaming animals exist in both the terrestrial environment (e.g. cougars [Dickson and Beier 2002], golden eagles [Domenech et al. 2015], and foxes [Chamberlain and Leopold 2000]) and the aquatic realm (e.g. catfish [Kadye and Booth 2012], octopuses [Regueira et al. 2013 and cardinalfishes [Marnane 2000]). Though these animals migrate frequently, they often have home bases and exhibit high site-fidelity. Insight into adult habitat selection at all scales is critical as the ecological baselines of numerous systems are rapidly shifting. This shift is projected to most devastatingly influence species that are tightly coupled with specific habitats (Munday 2004). Specialized habitat selection is particularly important in the intensely complex and diverse coral reef ecosystem (Huston 1985). The selection of favorable habitat can increase overall fitness through predator avoidance (Caley and St John 1996), food availability (Pereira et al. 2012), and access to mating opportunities (Baeza et al. 2016). Due to competition for space and the potential consequences of selecting inappropriate habitat (e.g., death), natural selection should favor individuals that choose suitable sites. On coral reefs, this selection has resulted in a high proportion of species exhibiting obligate habitat use (Connell 1978).

64 Obligate habitat associations often begin at the conclusion of the larval stage. Thereafter, the inhabitant rarely leaves the host, as seen in the anemonefish − sea anemone relationship (Dixson et al. 2014). However, some reef animals occupy specific habitats during rest periods, yet vacate to forage or mate. Cardinalfishes (Apogonidae), for example, reside in caves or dendritic corals during the day (Greenfield and Johnson 1990) but leave at night to forage throughout the reef (Chave 1978). Host specificity in cardinalfishes is pronounced, with individuals showing high site fidelity by homing back to specific locations within a coral matrix after being displaced 1-2 km (Marnane 2000). Such temporal homing has been observed in multiple species of reef teleosts (Ogden & Buckman 1973; Quinn and Brodeur 1991), as well as several invertebrate groups including crustaceans (Herrnkind and McLean 1971; Hahn and Itzkowitz 1986; Vannini and Cannicci 1995), cephalopods (Mather 1991), and limpets (Cook et al. 1969), yet the mechanisms behind the selection of habitat are relatively unknown. Shrimp associations within tube sponges are rare among the family,

Hippolytidae, which consists primarily of cleaner and peppermint shrimp (Rhyne and Lin 2006). The peppermint shrimp, Lysmata pederseni, exhibit obligatory associations with tube sponges, particularly the sponge genera Callyspongia, Niphates, and

Aplysina throughout Caribbean reefs (Rhyne and Lin 2006; Baeza 2010; Baeza et al. 2016). Although L. pederseni is not a true cleaner shrimp, it may passively clean its surroundings (Rhyne and Lin 2006), potentially contributing to sponge health. Further, these cathemeral shrimp are ecologically distinct from other peppermint shrimp due to their propensity for living alone or in small groups, rather than gregariously (Rhyne

65 and Lin 2006). Little is known about the external factors that influence the unique habitat association of L. pederseni within tube sponges. The evaluation of potential hosts can be accomplished through chemosensory perception or an assessment of morphological characteristics (Enright 1978). The relative use of these stimuli depends on species, spatial proximity, and environmental variables (Kingsford et al. 2002). Lysmata pederseni assess chemical cues using olfactory receptors in aesthetascs on their antennules (Hallberg et al. 1992). These shrimp are capable of detecting and responding to the chemical cues from the sponge Callyspongia vaginalis (Baeza et al. 2016); however the role chemical cues play in species-specific habitat selection has yet to be understood. Additionally, host morphology can influence habitat choice, as size and shape can be critical not only for protection from predators, but also for space required to perform basic functions such as mating (Vytopil and Willis 2001). Morphological assessment can be accomplished using visual and tactile cues, making it a reliable source of information even for visually-limited shrimp (Caves et al. 2016).

In this study, we investigated the tube sponge habitat selection of adult L. pederseni. First, the movement patterns of adult peppermint shrimp were identified using a mark-and-recapture experiment. Then, belt transects were conducted to investigate natural variations in shrimp-associated sponge species and morphologies. Subsequently, to determine which sensory cues are pertinent for habitat selection, both chemical and morphological preferences were tested in isolation. Finally, as the natural distribution of sponge species varies spatially, patch reefs were constructed to test sponge preference when all cues were present, and sponges were equidistant. Habitat specialists are more susceptible to the shifting species composition on coral

66 reefs, especially when associated with living hosts. A better understanding of vital processes, such as specialized habitat selection, can lead to more thorough predictions of how ecosystems and communities will respond to future conditions. We use L. pederseni as a model organism in this study, but the results have the potential to inform habitat selection across diverse taxa.

3.3 Methods

3.3.1 Temporal movements

Lysmata pederseni habitat fidelity was monitored during the day and night to gain insight into the extent and temporal scale of adult movement and habitat selection. Short-term habitat fidelity was assessed using an overnight mark-and- recapture technique at two reefs near Carrie Bow Cay, Belize (16°48'9.26"N, 88° 4'54.87"W; fore reef: 17 m max depth, surveyed August 23-25, 2016; lagoon reef: 9 m max depth, surveyed June 17-18 and 24-27, 2017). Sponges containing L. pederseni were marked with flagging tape in the afternoon (~16:00). If multiple sponge tubes hosted shrimp, each tube was individually flagged. Each shrimp within the sponge was carefully extracted by squeezing the sponge so that the shrimp rose through the column into a net (Baeza et al. 2016) and tagged using an elastomer tag (Northwest

Marine Technology, Inc., USA) on the ventral side of the sixth abdominal segment (Baeza 2010). After tagging, shrimp were returned to their original sponge tube. All shrimp were alive after release (n=26). Shrimp locations were assessed three times following tagging. First, presence of shrimp was recorded the morning after the tagging procedure (~10:00) to appraise site fidelity. The subsequent night, shrimp-sponge associations were evaluated using a

67 1000 lumen flashlight at ~21:00 to determine whether shrimp nocturnally vacate their host sponge. Shrimp could be detected in a few seconds, which minimized disturbance by dive lights. Finally, flagged sponges were checked for shrimp presence the morning after the nocturnal observations (~10:00) to confirm site fidelity despite disturbance.

3.3.2 Natural habitat associations Natural variability in shrimp-sponge associations was determined using 15x1 m belt transects on reefs around Carrie Bow Cay, Belize between March 31-April 2, 2016 (depth range: 3-14 m, n=22). Along each transect, all tube sponges were inspected to quantify abundance of resident L. pederseni and four sponge parameters were recorded: (1) sponge species, (2) number of tubes, (3) tube height from base to tip, and (4) osculum diameter, measured at the widest point.

3.3.3 Morphological preference

Shrimp used in the cue isolation and patch reef experiments were collected using methods described above and transported to the Smithsonian Research Station on Carrie Bow Cay, where they were held in ~15 L flow-through aquaria with coral rubble for shelter until testing. Cafeteria-style choice experiments were used to assess shrimp morphological preference to host sponges. Four morphological types of artificial tube sponges were constructed from black foam pool noodles (Fig. 3.1): tall wide (20 cm high, 7 cm osculum diameter), short wide (10 cm high, 7 cm osculum diameter), tall narrow (20 cm high, 3 cm osculum diameter), and short narrow (10 cm high, 3 cm osculum diameter). One of each sponge morphotype was fixed in a random configuration to an acrylic plate such that each artificial sponge was 30 cm from the center. The acrylic plates with attached artificial sponges were placed into circular

68 aquaria (40L) and weighed down with dive weights. Each aquarium was filled with seawater and sand was added to eliminate all crevices. A 10 cm diameter habituation chamber, constructed from plastic mesh (1.27 x 1.27 cm) and fly screen (1x1 mm), was positioned upright in the sand such that the top of the chamber was above the waterline. Each aquarium contained an airstone. The sand and water were replaced after each trial. After sunset (~21:00), one shrimp was carefully placed inside the habituation chamber for 10 min. At the end of the habituation period, the chamber was slowly removed, and the aquarium was covered with a mesh screen. This allowed natural light to penetrate but prevented the shrimp from escaping. After 12 hours, the shrimp location was recorded (n=36).

69 22 tall, narrow shrimp selected3 cm tubes 8 1.5 cm tall, wide shrimp selected cm 20 0 7 cm tubes short, wide shrimp selected 1.5 cm 7 cm 30.48 cm 30.48 tubes

1.5 cm

20 cm 20 10 cm 10 3 cm

1.5 cm

10 cm 10 0 short, narrow shrimp selected 38.1 cm tubes Figure 3.1: Infogram of the experimental methods to test sponge morphology preferences in isolation. Grey cylinders represent artificial sponges made of foam pool noodles. Experimental results are shown in text boxes with a significant preference for tall and narrow tube morphology.

3.3.4 Chemical preference To assess the use of chemical cues in habitat selection, shrimp preference was tested using an Atema two-channel choice flume (Gerlach et al. 2007). The choice flume (23.5 x 4 cm) allows an individual organism to experience the chemical cues of two different water sources simultaneously by presenting them side-by-side. The two water sources were gravity-fed from buckets through tubes into the choice chamber. Laminar flow was maintained at 100 mL min-1 using flow meters (Dwyer MMA-40) and checked periodically using dye tests. A shrimp was placed in the center of the flume and given a 2 min habituation period, during which time the shrimp was free to move throughout the chamber and explore either cue. After the habituation period, the

70 shrimp’s position in each stimulus was recorded at 5-second intervals for 2 min. The water sources were then switched to eliminate a potential side bias with a 2-min flushing period and the entire test was repeated. Each shrimp was tested only once per trial (n=10) and all trials were run blind. Sponge species used as cues in fluming trials were chosen based on habitat associations observed during the transects (Table 3.1). Lysmata pederseni associated with the tube sponges, Callyspongia vaginalis, C. plicifera and Niphates digitalis. In contrast, another common tube sponge, Aplysinia fistularis, never hosted peppermint shrimp and was used as a negative control cue. Sponges used for cue generation were cut at the base of their structures and transported to the lab. Only sponges lacking resident shrimp were collected and any additional epibionts were removed. To generate chemical cues, each species of sponge was spun 20 times in a salad spinner to eliminate excess water before being weighed to 20 g and added to 2 L of reef water for 1 hour in a closed system. This solution was diluted to 10 L using reef water. Water collected directly from the reef acted as a general reef water cue. Chemical preferences were analyzed using the Kolmogorov-Smirnov (K-S) nonparametric test. The proportion of time spent in each cue was compared to the proportion of time spent on one side of the choice flume when no cues were present (blank control).

71 Table 3.1: Chemical choice comparisons. * indicates non-associated habitat based on transect data.

Choice trials Preference hierarchy trials

A. fistularis vs Reef water C. vaginalis vs A. fistularis*

C. vaginalis vs Reef water C. vaginalis vs C. plicifera

C. plicifera vs Reef water C. vaginalis vs N. digitalis

N. digitalis vs Reef water C. plicifera vs N. digitalis

Reef water vs Reef water Reef water vs Reef water

3.3.5 Patch reefs

The patch reef experiment was conducted in a large sand patch (8m depth, 16°48'45.15"N, 88° 5'8.29"W) to test the preference for different sponge species when all cues were present and sponge distribution was equal (Fig. 3.2). Eight patch reefs were constructed, and each contained one sponge of each of the four test species, C. vaginalis, C. plicifera, N. digitalis and A. fistularis. Sponges were arranged randomly and size matched based on height and overall mass, as mass can influence the quantity of chemical cues released. Number of tubes and tube width were distinct morphological characteristics typical of specific sponge species and were therefore not size matched. Two rubble pieces were added to the center of the patch and each sponge was 0.5 m from this point and equidistance from each other. Experimental shrimp were tagged with an elastomer tag to indicate the sponge species from which it was collected. Shrimp were placed on patch reefs at ~21:00 (n=36). Dive lights never directly illuminated the patch reefs. At each patch, a 10 cm habituation chamber was wedged into the sand between the rubble pieces and one shrimp was carefully placed within this chamber. The chamber was open at the top,

72 allowing the shrimp to leave and explore. After 30 min, the habituation chambers were removed. If the shrimp was still inside, the sides of the chamber were gently squeezed until the shrimp crawled onto the rubble pieces. After a 12-hour overnight period, shrimp location was recorded, and shrimp were removed from the patch reef. The sponges were shifted one position clockwise each morning to eliminate potential biases due to current direction or light availability. The experiment was conducted over six nights using different shrimp each time.

16 C. vaginalis 8 shrimp selected Habituation A. Pistularis chamber shrimp selected

0.5 m rubble 0.5 m 5 2 C. plicifera N. digitalis shrimp selected 0.5 m shrimp selected 0.5 m

Rotated one position clockwise each day Figure 3.2: Infogram of the experimental methods to test sponge preference when all cues were present, and sponges were equidistant. Experimental results are shown in text boxes with a significant preference for C. vaginalis.

73 3.4 Results

3.4.1 Temporal movements

Morning observations following afternoon tagging revealed that 96.2% ± 7.3 of tagged shrimp were found in the same sponge individual, and only 8.0% ± 10.6 of those found were within a different tube in the same sponge. Nocturnal observations conducted the subsequent night demonstrated definite movement from host sponges; 80.8% ± 15.1 of tagged shrimp vacated their host sponge entirely, with an additional 11.5% ± 12.3 migrating between different tubes of the same sponge. The majority of the tagged shrimp were absent from within or around the sponge individual; however, five tagged shrimp were observed outside of their host sponge tubes over the course of the experiment. Multiple untagged shrimp that were not present during the day were observed in flagged sponges at night. The morning following the nocturnal observations, 80.8% ± 15.1 of the tagged shrimp were found in the same sponge that they were originally tagged in, and only 23.8% ± 18.2 of those were found in a different tube within the same sponge. The proportion of shrimp present in their host sponges during the day compared to the night was statistically different (z-test for proportions in R [R Core Team 2017]; p<0.0001).

3.4.2 Natural habitat associations Transects confirmed the natural association of L. pederseni to specific tube sponge species (See Supplementary Data). The number of tubes per sponge significantly influenced shrimp occupancy rates (ANOVA [JMP Pro 13]; F

7,148=56.956, p<0.0001) and among naturally shrimp-associated sponges, both sponge species (ANOVA; F2,102=4.8863, p=0.0094) and number of tubes (ANOVA;

F6,102=99.64, p<0.0001) affected shrimp presence. The number of tubes, however, was

74 highly correlated to the sponge species (ANOVA; F9,148=7.056; p<0.0001). When sponge tube morphology was analyzed independent of sponge species, L. pederseni were more abundant on sponges with high height to diameter ratios (Wilcoxon signed- rank test [JMP Pro 13]; p=0.0477, n=234), thereby favoring tall, narrow sponges opposed to short, wide sponges (Fig. 3.3). An analysis of individual species morphologies reveals that L. pederseni tend to occupy N. digitalis tubes with high height to diameter ratios (Wilcoxon signed-rank test; p=0.0012, n=89). No morphological inclinations were observed for C. vaginalis (Wilcoxon signed-rank test; p=0.1605, n=16) or C. plicifera (Wilcoxon signed-rank test; p=0.4768, n=42). The natural abundance of sponge species varies spatially; when the relative population abundance of each sponge species was considered, a greater percentage of C. vaginalis tubes hosted L. pederseni (18.8%, n=16), followed by N. digitalis (8.9%, n=89) and lastly C. plicifera (4.8%, n=42). Additionally, tube sponges hosted other epibionts including fish, crabs, brittle stars, and non-conspecific shrimp, but the presence of other inhabitants did not preclude the focal shrimp from association with sponge individuals.

75 0.8

0.7

0.6 0.5 * *

Shrimp habitat 0.4

0.3

0.2

Ratio of sponge height/diameter (log) Ratio of sponge height/diameter

Shrimp habitat No Shrimp Shrimp habitat No Shrimp No Shrimp Shrimp habitat No Shrimp 0.1

0 C. vaginalis C. plicifera N. digitalis

All sponges Figure 3.3: Natural shrimp-sponge associations. Transect data reveals a preference for tall narrow sponge tubes when all sponges are combined and when comparing C. vaginalis alone. White bars represent sponge tubes that did not host shrimp while grey bars represent sponge tubes with resident shrimp. * indicates significance.

3.4.3 Morphological preference When morphological preferences were tested in isolation using artificial sponges, no shrimp were found in association with short sponges, regardless of osculum width (Fig. 3.1). Shrimp significantly preferred taller (χ2 goodness-of-fit test in [R Core Team 2017]; χ2= 28.033, p<0.0001) and narrower (χ2 goodness-of-fit test; χ2= 5.633, p=0.0176) tubes. Of the 36 shrimp tested, 6 did not choose an artificial sponge, 22 selected tall, narrow sponges and 8 selected tall, wide sponges.

3.4.4 Chemical preference

Shrimp preferred the chemical cues of C. vaginalis, C. plicifera and N. digitalis to the general reef cue, by spending >80% of their time in the side of the

76 flume containing the sponge cue (K-S test [JMP Pro 13]; p<0.0001). However, L. pederseni avoided the chemical cue from the negative control, A. fistularis, only spending 21% ± 2.66 SE of the time in this cue (K-S test, p<0.0001) (Fig. 3.4a). When presented with two different sponge cues simultaneously, peppermint shrimp preferred C. vaginalis to all other species tested (K-S test; p<0.0001 for either comparison). When chemical cues from C. plicifera were tested against the chemical cues of N. digitalis, shrimp preferred C. plicifera (K-S test; p<0.0001) (Fig. 3.4b).

77 a 100 * * * 80 *

2 O

40 2

Sponge Cue Sponge

Reef H Reef

O 2

20 Reef H

Reef H Reef

Sponge Cue Sponge

Sponge Cue Sponge Sponge Cue Sponge Reef H Reef 0

A. Pistularis N. digitalis C. plicifera C. vaginalis b 100 ** *

80 * Percent time in cue in time Percent

C. plicifera C.

istularis P

20 A.

N. digitalis N.

N. digitalis N. C. vaginalis C. C. plicifera C. C. vaginalis C. C. vaginalis C. 0

Choice trials Figure 3.4: Chemical choice trials. (A) Shrimp response to individual sponge cues compared to general reef cues. (B) Shrimp response when presented with two sponge cues simultaneously to determine a hierarchical preference. All chemical data are mean percent time ± standard error. * indicates significance.

78 3.4.5 Patch reefs The sponge species that shrimp were collected from did not impact sponge choice in the patch reef experiment (Fisher’s exact test in R [34]; p=0.3512), allowing data from all of the shrimp to be pooled. Sponge selection by L. pederseni was not random (χ2 goodness-of-fit test in [R Core Team 2017]; χ2=14.032, p=0.002862); rather, L. pederseni significantly preferred C. vaginalis, with 51% (n=16) of shrimp associating with this species the following morning. C. plificera was selected by 16% (n=5) of the shrimp and 6% (n=2) of the shrimp selected N. digitalis (Fig. 3.2). Surprisingly, 26% of the shrimp (n=8) selected A. fistularis, an association never observed naturally. Five shrimp either abandoned the patch reef or succumbed to predation.

3.5 Discussion

As the shifting climate causes changes in local species abundances (Hughes et al. 2003), habitat specialists will be some of the most vulnerable due to their association with living hosts (Munday 2004). Understanding specialized habitat associations and the mechanisms used in assessing host individuals is vital at both the larval and adult stages. Here, we show that adult L. pederseni vacate and re-select habitat daily, with strong site fidelity towards the same host individual. These shrimp demonstrate a preference for the chemical cues of specific sponge species as well as the morphological characteristics of tall narrow sponge tubes, a pattern that is reflected in both natural benthic transects and the in situ patch reef experiment. The natural distribution of sponges, however, can also influence sponge-shrimp associations across the reef, thereby creating high variation in chosen host sponges.

79 Most obligate habitat associations begin at the larval stage and thereafter the inhabitant remains solely within the host. In contrast, L. pederseni show high site fidelity to individual sponges, yet vacate at night, suggesting that adult habitat re- selection is frequent. Previous studies have indicated that L. pederseni host fidelity can be constant for up to two months (Baeza 2010), depending on the sexual phase or mating system (Baeza et al. 2016). Lysmata pederseni are protandric simultaneous hermaphrodites, where all juveniles are male and develop into functional hermaphrodites (Baeza 2009). Populations can tend towards monogamy or polygynandry depending on location. Populations at Carrie Bow Cay, Belize are primarily monogamous (Baeza et al. 2016), whereas populations in the Florida Keys exist in polygynandrous relationships characterized by promiscuity and frequent host switching (Baeza 2010). Previous studies, however, have only assessed site fidelity during the day. Our study confirms that regular nocturnal vacancies occur despite apparent daytime fidelity to specific sponges, suggesting that habitat selection is an essential process at the adult stage even among monogamous populations.

Morphological characteristics tested in isolation revealed a preference for tall, narrow sponges. This morphological affiliation is confirmed in field transects, but whether these natural associations are due to shrimp selection or differential survival remains to be determined. Specifically, morphological characteristics of C. vaginalis and C. plicifera do not influence the likelihood of shrimp association, possibly due to their ubiquitous tube-like shapes. Conversely, shrimp are more abundant on tall, narrow morphotypes of N. digitalis. Tube shape of N. digitalis varies with some having an irregularly shaped osculum that is long and narrow. Measurements were taken at the widest diameter of the osculum, causing irregular morphotypes to have

80 low height to diameter ratios. Wider openings may allow easier predator access to the epifaunal community within and may promote shrimp to avoid this morphotype. Height preference in this species is further confirmed in the Florida Keys where L. pederseni were never found in sponges smaller than 10 cm high (Baeza et al. 2016). While our isolation experiment validates this, one shrimp was found naturally within a 6 cm C. plicifera tube. Lysmata pederseni easily discriminated between different sponge odours. Chemical preferences (C. vaginalis, C. plicifera and N. digitalis) and chemical deterrents (A. fistularis) matched natural presence-absence field observations. Previous work on the chemical detection of L. pederseni found that these shrimp positively respond to the chemical cues of C. vaginalis (Baeza et al. 2016). Our study corroborates and expands on this research by testing the response to additional host species and identifying hierarchical preferences. Lysmata pederseni have the strongest preference for the chemical cues of C. vaginalis, followed by C. plicifera and N. digitalis. This ranking is supported by previous field studies stating that L. pederseni are more commonly or only found in C. vaginalis (Rhyne and Lin 2006; Baeza 2010; Baeza et al. 2016). Reef sponges can be classified into 3 categories in relation to chemical defenses and predation rates: (1) chemically-defended species, (2) chemically- undefended species that persist because of high rates of growth, reproduction or healing, and (3) chemically-undefended species that persist in secluded refuges (Pawlik 2011). Callyspongia and Niphates are fast-growing and chemically undefended (Pawlik et al. 1995), whereas Aplysina sponges produce secondary metabolites that defend against predation (Pawlik et al. 1995), fouling (Willemsen

81 1994), microbial growth (Kelly et al. 2005), and allelopathic attacks (Pawlik et al. 2007). Aplysina fistularis, in particular, naturally extrudes secondary metabolites, but the rate of extrusion can increase when damaged (Walker et al. 1985). Transect and fluming data indicate that L. pederseni prefer the fast-growing, chemically undefended species, suggesting that sponge morphology may play a larger role in shrimp preference than the chemical defense of the habitat itself. The differential distribution of sponges across the reef could be a major factor contributing to sponge-shrimp associations. Transects revealed more L. pederseni residing in N. digitalis than other species; however, after factoring in sponge population sizes, peppermint shrimp inhabited a higher proportion of C. vaginalis sponges, matching the chemical preference data. When the influence of distance and scarcity was eliminated in the patch reef experiment, L. pederseni preferred C. vaginalis, regardless of origin sponge. Although natural sponge associations and chemical cue preferences corroborate one another, the patch reef data demonstrated the willingness of shrimp to associate with A. fistularis. This sponge never hosted a shrimp naturally and was avoided using chemical cues alone. The association of eight L. pederseni individuals with A. fistularis in the patch reefs indicates that the secondary metabolites produced by this sponge are not acutely toxic to the shrimp; however, as seen with other sponge dwelling species, these secondary metabolites could have chronic effects (Henkel and Pawlik 2011). The growth rate of the brittle star, Ophiothrix lineata, was significantly reduced when experimentally forced to reside in a sponge species that produces secondary metabolites rather than in a preferred non-defended sponge (Henkel & Pawlik 2011). The selection of A. fistularis by peppermint shrimp in the patch reef also supports the idea that sponge morphology

82 has a greater influence than sponge chemistry on short-term habitat selection. Aplysina fistularis had the highest height to diameter ratio of all tube sponges on the transects, potentially making this species a beneficial choice for a shrimp looking to quickly escape from predators. The absence of L. pederseni in A. fistularis sponges in nature suggests that despite beneficial morphology, the secondary metabolites produced by this sponge may have long-term consequences. In conclusion, L. pederseni utilize morphology as an effective habitat selection tool and also show distinct chemical preferences for specific hosts. Habitat selection in adults is understudied compared to habitat selection in larvae but is no less important as movements between distinct habitats can influence material transport, nutrient fluxes and community dynamics (Marnane 2000). Understanding how and why adult habitat specialists choose their particular habitat is important in today’s rapidly changing world. Habitat degradation is occurring at an unprecedented rate, especially in reef environments (De’ath et al. 2012). Habitat specialists may be more threatened by environmental change when they are associated with a living host because the host can also be impacted by changing conditions (Munday 2004). Since habitat specialists can drive biodiversity (Sale 1977), an understanding of the mechanisms behind habitat selection can inform research and management practices.

3.6 Acknowledgements I would like to acknowledge R. Brooker and D. Dixson for contributions to experimental design and data collection. I would also like to thank S. Carlson, Z. Cowan, Z. Foltz, L. Johnston, S. Jones, J. Joseph, A. Looby, V. Paul and J. Sneed, for helping with data collection and A. Franca and J. Biddle for help with sponge identification.

83 REFERENCES

Baeza, JA. 2009. Protandric simultaneous hermaphroditism is a conserved trait in Lysmata (Caridea: Lysmatidae): implications for the evolution of hermaphroditism in the genus. Smithson Contrib Mar Sci 38:95–110.

Baeza JA. 2010. The symbiotic lifestyle and its evolutionary consequences: social monogamy and sex allocation in the hermaphroditic shrimp Lysmata pederseni. Naturwissenschaften. 97:729-741.

Baeza JA, Guéron R, Simpson L, Ambrosio LJ. 2016. Population distribution, host- switching, and chemical sensing in the symbiotic shrimp Lysmata pederseni: implications for its mating system in a changing reef seascape. Coral Reefs. 35:1213-1224.

Caley MJ, St John J. 1996. Refuge availability structures assemblages of tropical reef fishes. J Anim Ecol. 65:414-428.

Caves EM, Frank TM, Johnsen S. 2016. Spectral sensitivity, spatial resolution and temporal resolution and their implications for conspecific signaling in cleaner shrimp. J Exp Biol. 219:597-608.

Chamberlain MJ, Leopold BD. 2000. Spatial use patterns, seasonal habitat selection, and interactions among grey foxes in Mississippi. J Wildl Manage. 64:742- 751.

Chave EH. 1978. General ecology of six species of Hawaiian cardinalfishes. Pac Sci. 32:245–270.

Connell JH. 1978. Diversity in tropical rain forests and coral reefs. Science.199:1302- 1310.

Cook A, Bamford OS, Freeman JDB, Teideman DJ. 1969. A study of the homing habit of the limpet. Anim Behav. 17:330-339.

Cowen RK, Sponaugle S. 2009. Larval dispersal and marine population connectivity. Ann. Rev. Mar. Sci. 1:443-466.

84 De’ath G, Fabricius KE, Sweatman H, Puotinen M. 2012. The 27–year decline of coral cover on the Great Barrier Reef and its causes. Proc Natl Acad Sci U.S.A. 109:17995-17999.

Dickson BG, Beier P. 2002. Home-range and habitat selection by adult cougars in Southern California. J Wildl Manage. 66:1235-1245.

Dixson DL, Jones GP, Munday PL, Planes S, Pratchett MS, Thorrold SR. 2014. Experimental evaluation of imprinting and the role innate preference plays in habitat selection in a coral reef fish. Oecologia 174:99.

Domenech R, Bedrosian BE, Crandall RH, Slabe VA. 2015. Space use and habitat election by adult migrant golden eagles wintering in the Western United States. J Raptor Res. 49:429-440.

Enright JT. 1978. Migration and Homing of Marine Invertebrates: A Potpourri of Strategies. In Schmidt-Koenig K, Keeton WT, editors. Animal Migration, Navigation, and Homing. Berlin, Heidelberg: Springer. p. 440-446

Gerlach G, Atema J, Kingsford MJ, Black KP, Miller-Sims V. 2007. Smelling home can prevent dispersal of reef fish larvae. Proc Natl Acad Sci U.S.A. 104:858- 863.

Greenfield DW, Johnson RK. 1990. Heterogeneity in habitat choice in cardinalfish community structure. Copeia. 1107–1114.

Hahn P, Itzkowitz M. 1986. Site preference and homing behavior in the mysid shrimp Mysidium gracile (Dana). Crustaceana. 51:215-219.

Henkel TP, Pawlik JR. 2011. Host specialization of an obligate sponge-dwelling brittlestar. Aquat Biol. 12:37-46.

Herrnkind WF, McLean R. 1971. Field studies of homing, mass emigration, and orientation in the spiny lobster, Panulirus argus. Ann NY Acad Sci. 188:359- 376.

Hallberg E, Johansson KU, Elofsson R. 1992. The aesthetasc concept: structural variations of putative olfactory receptor cell complexes in Crustacea. Microsc Res Tech. 22:325-335.

Hughes TP, Baird AH, Bellwood DR, Card M, Connolly SR, Folke C, et al. 2003. Climate change, human impacts, and the resilience of coral reefs. Science. 301:929-933.

Huston MA. 1985. Patterns of species diversity on coral reefs. Annu Rev Ecol Evol Syst. 16:149-177.

85 JMP Pro, Version 13, SAS Institute Inc., Cary, NC, 1989-2007.

Kadye WT, Booth AJ. 2012. Movement patterns and habitat selection of invasive African sharptooth catfish. J Zool. 289:41-51.

Kelly SR, Garo E, Jensen PR, Fenical W, Pawlik JR. 2005. Effects of Caribbean sponge secondary metabolites on bacterial surface colonization. Aquat Microb Ecol. 40:191-203.

Kingsford MJ, Leis JM, Shanks A, Lindeman KC, Morgan SG, Pineda J. 2002. Sensory environments, larval abilities and local self-recruitment. Bull Mar Sci. 70:309-340.

Marnane MJ. 2000. Site fidelity and homing behaviour in coral reef cardinalfishes. J Fish Biol. 57:1590-1600.

Mather JA. 1991. Navigation by spatial memory and use of visual landmarks in octopuses. J Comp Physiol A Neuroethol Sens Neural Behav Physiol. 168:491- 497.

Meadows PS, Campbell JI. 1972. Habitat selection by aquatic invertebrates. Adv Mar Biol. 10:271-382.

Munday PL. 2004. Habitat loss, resource specialization, and extinction on coral reefs. Glob Change Biol. 10:1642-1647.

Ogden JC, Buckman NS. 1973. Movements, foraging groups, and diurnal migrations of the striped parrotfish Scarus croicensis Bloch (Scaridae). Ecology. 54:589– 596.

Pawlik JR. 2011. The chemical ecology of sponges on Caribbean reefs: natural products shape natural systems. BioSci. 61:888–898.

Pawlik JR, Chanas B, Toonen RJ, Fenical W. 1995. Defenses of Caribbean sponges against predatory reef fish. I. Chemical deterrency. Mar Ecol Prog Ser. 127:183-194.

Pawlik JR, Steindler L, Henkel TP, Beer S, Ilan M. 2007. Chemical warfare on coral reefs: Sponge metabolites differentially affect coral symbiosis in situ. Limnol. Oceanogr. 52:907-911.

Pereira PHC, Leal ICS, de Araújo ME, Souza AT. 2012. Feeding association between reef fishes and the fire coral Millepora spp. (Cnidaria: Hydrozoa). Mar Biodivers Rec. 5:e42.

86 Quinn TP, Brodeur RD. 1991. Intra-specific variations in the movement patterns of marine animals. Amer Zool. 31:231-241.

R Core Team. 2017. R: A language and environment for statistical computing. R Foundation for Statistical Computing, Vienna, Austria. URL https://www.R- project.org/.

Regueira M, González AF, Guerra A. 2013. Habitat selection and population spreading of the horned octopus Eledone cirrhosa (Lamarck, 1798) in Galician waters (NW Atlantic). Fish Res. 152:66-73.

Rhyne AL, Lin J. 2006. A western Atlantic peppermint shrimp complex: redescription of Lysmata wurdemanni, description of four new species, and remarks on Lysmata rathbunae (Crustacea: Decapoda: Hippolytidae). Bull Mar Sci. 79:165-204.

Sale PF. 1977. Maintenance of high diversity in coral reef fish communities. The Am Nat. 111:337-359.

Vannini M, Cannicci S. 1995. Homing behaviour and possible cognitive maps in crustacean decapods. J Exp Mar Biol Ecol. 193:67-91.

Vytopil E, Willis B. 2001. Epifaunal community structure in Acropora spp.(Scleractinia) on the Great Barrier Reef: implications of coral morphology and habitat complexity. Coral Reefs. 20:281-288.

Walker RP, Thompson JE, Faulkner DJ. 1985. Exudation of biologically-active metabolites in the sponge Aplysina fistularis. Mar Biol 88:27-32.

Willemsen PR. 1994. The screening of sponge extracts for antifouling activity using a bioassay with laboratory-reared cyprid larvae of the barnacle Balanus Amphitrite. Int Biodeterior Biodegradation. 34:361-373.

87 Chapter 4

CORAL MORPHOLOGY IS LINKED TO INCREASED ABUNDANCE AND FITNESS OF OBLIGATE CORAL-DWELLING SHRIMP, CORALLIOCARIS GRAMINEA.

4.1 Abstract

Branching corals act as hosts to a wide variety of epibionts, many of which are obligate associates that depend on corals for survival. Appropriate hosts can increase the fitness of inhabitants through greater access to resources. However, the suitability of a host can vary based on species and morphological characteristics. This study investigates the influence of coral species and morphology on the fitness and abundance of the obligate coral-dwelling shrimp, Coralliocaris graminea. Transects were conducted at two locations in the Fijian Islands and morphological parameters were recorded from each corymbose or digitate acroporid coral intersected. Resident C. graminea were removed from the corals to assess body size, claw size, and fecundity. Shrimp associated with 7 species of Acropora corals, and the distribution of shrimp within these corals was not random. Coral morphology was a strong indicator of shrimp association; C. graminea tended to inhabit corals with larger diameters and higher branch heights. Fitness of C. graminea, quantified by body size, claw size, and fecundity, was higher for those inhabiting larger, deeper corals and corals that hosted conspecifics. Movement of C. graminea between corals has not been documented but must be occurring, evidenced by the presence of gravid females residing in corals without males present. This study suggests that coral preference of C. graminea is not species specific and coral morphology may have a greater influence on shrimp fitness

88 and abundance; therefore, these obligate coral inhabitants may survive on structurally similar, non-host corals. As shrimp abundance and fitness increased with coral head diameter, conservation efforts should aim to increase individual colony size along with overall coral coverage.

4.2 Introduction Coral reefs have declined rapidly in the last few decades in both the Caribbean (Hughes and Tanner 2000) and the Indo-Pacific (Bruno and Selig 2007; Hughes et al. 2018). This deterioration is characterized by reduced coral coverage (Gardner et al. 2003; Bruno and Selig 2007) and phase shifts (Hughes et al. 2007), which lead to dwindling fish abundance and diversity (Jones et al. 2004), local extinctions (Munday et al. 2004), and decreased resilience (Anthony et al. 2011). Differences in life history, evolutionary experiences, and genetic diversity have resulted in variable tolerance to stress among corals species (Hughes et al. 2003; Pandolfi et al. 2011). This tolerance variation may allow coral reefs as an ecosystem to withstand environmental change, but sensitive species, as well as coral obligate species, will face the threat of extinction. For example, the vital reef-building genus, Acropora, is among the most sensitive to environmental change (Loya et al. 2001; Comeau et al. 2014) and has already experienced massive declines worldwide (Williams and Miller 2012). The loss of high-complexity corals, such as Acropora spp., is especially devastating as they provide structure and substrate for numerous organisms (Jones et al. 2004; Wild et al. 2011). Large, structurally complex corals host numerous obligate associates that depend on the coral for survival and are invariably found in association with their hosts (Stella et al. 2010). These obligate inhabitants acquire food resources while in

89 their habitat and these resources are allocated to various functions including tissue maintenance, reproduction, defense, and storage (Cronin 2001). In a resource-limited environment, acquired energy must be distributed between these functions in a way that provides the best chance of survival. However, when resources abound, more energy can be allocated to more functions. The availability of resources can be highly correlated to habitat (MacArthur and Levins 1964), especially for obligate inhabitants (Munday 2004). A suitable habitat will allow residents to acquire more resources, thereby increasing individual growth, fecundity, and overall fitness. The relationship between coral morphology and epibiont fitness is investigated using the obligate coral-dwelling shrimp, Coralliocaris graminea. This cryptic snapping shrimp is found within numerous species of corymbose (bushy) or digitate corals throughout the Indo-Pacific (Bruce 1998). While these shrimp typically associate exclusively with acroporid corals, rare individuals have been found within Stylophora, Seriatopora and Pocillopora (Miyake 1968; Bruce 1972, 1977). Shrimp within these alternative genera were characterized as small juveniles, indicating that these corals may not be ideal habitat but rather a transitional host (Bruce 1977). Among the acroporids, C. graminea has been observed associating with at least 18 species (Miyaki 1968; Bruce 1972,1976, 1977; Patton 1994; Stella et al. 2010; Vytopil and Willis 2011). This study seeks to understand the influence of coral species and morphology on the abundance and condition of C. graminea. Using extensive transects at two Fijian locations, the percent occupancy of the focal shrimp within Acropora corals was quantified. Body size, claw size and fecundity were used as proxies for general fitness of the individuals. Studies involving C. graminea are rare in established literature and

90 none to date focus explicitly on the habitat preferences and associated fitness of this cryptic species. Declining reefs impact habitat specialized organisms disproportionately; therefore, investigating the full impact of coral coverage and morphology on the habitat associations of selective organisms can inform our understanding of future reef ecosystems.

4.3 Methods Forty-five 25 m transects assessing shrimp-coral associations were conducted along the reefs of Tavewa and Kadavu Islands, Fiji. Twenty transects were run on the southern coast of Kadavu Island (19° 2'46.53"S, 178°23'57.75"E) from 17-24 October 2016; an additional twenty-five transects were conducted on the eastern shore of Tavewa Island (16°55'23.32"S, 177°22'1.40"E) from 13-17 July 2017 and 24 September 2018. In addition to transects, in order to increase the power of the analysis, corals throughout the uninspected reef were haphazardly sampled and searched for presence of C. graminea and only corals that housed the focal shrimp were recorded. At each corymbose or digitate Acropora coral intersected by the transect (n=462), coral parameters were recorded, including: 1) coral species, 2) diameter, measured at the widest point, 3) height of branches, measured from the deepest point, and 4) the number of resident C. graminea. In addition, each coral was photographed from above to measure distance between branch tips (inter-brachial width). Mean inter-brachial spacing was analyzed using ImageJ software by averaging the distance (mm) between two adjacent axial tips from five haphazardly selected points per coral head.

91 C. graminea observed within Acropora corals at Tavewa Island were extracted from their host corals using a clove oil, ethanol mixture (Javahery et al. 2012). Shrimp (n=69) were euthanized and preserved in 4% formaldehyde before being transferred to a 60% ethanol solution. Eggs were extracted from fecund shrimp (n=35) by carefully massaging the pleopod area below the abdomen with blunt forceps and counted under a dissection microscope (Stella et al. 2011). To assess impact of coral morphology on shrimp size, C. graminea total length (TL), standard length (SL), claw length and width, and carapace width were measured from images analyzed in ImageJ (Schindelin et al. 2012) (Figure 4.1). Additionally, observations of community ecology were recorded including sex of collected shrimp, sex ratios of populated corals, and rate of limb loss. Claw dimensions were only recorded for the largest fully- grown claw because numerous shrimp were experiencing various stages of claw regeneration in at least one claw.

92 0.25 cm

Claw Length

Claw Width

Carapace Width SL

Fecundity

Eggs TL

0.25 cm Figure 4.1: Focal shrimp, Coralliocaris graminea, depicting body (Standard length (SL), Total length (TL), Carapace width) and claw (length, width) measurements. Inlay: Posterior ventral view of gravid female C. graminea.

93 4.4 Results Fifteen species of corymbose or digitate Acropora corals were identified during transects; C. graminea were associated with seven of these species: Acropora divaricata, A. gemmifera, A. humilis, A. millepora, A. nasuta, A. tenuis and A. valida, henceforth known as preferred corals. Of the preferred corals, the majority of shrimp were associated with A. nasuta (57.5%), A. tenuis (15%), and A. millepora (14.2%); however, these species were among the most common along the transects. Therefore, coral species abundance was incorporated into the expected frequencies of a χ2 goodness-of-fit test (R, R Core Team 2017) to assess the role of coral species on shrimp abundance (Table 4.1). Observed shrimp-coral associations deviated significantly from what would be expected from a random distribution of shrimp (χ2=32.442, p<0.001). An assessment of standardized residuals indicated that the abundance of shrimp within A. humilis, A. nasuta, A. tenuis, and A. valida is higher than expected and the abundance within A. divericata, A. gemmifera, and A. millepora is lower than expected (Table 4.1).

94 Table 4.1: Observed and Expected shrimp abundance by coral species used in the χ2 goodness-of-fit test (χ2=32.442, p<0.001). The abundance of each coral species along the transect (column 2) was incorporated into the expected frequency of shrimp associations. Standardized Residuals (Std. Res.) indicate the extent and direction of deviation from expected values.

Coral species Coral abundance Observed shrimp Expected shrimp Std. Res A. divaricata 9 1 2.58 -0.98 A. gemmifera 61 3 17.47 -3.46 A. humilis 12 8 3.44 2.46 A. millepora 81 17 23.20 -1.29 A. nasuta 216 69 61.86 0.91 A. tenuis 35 18 10.02 2.52 A. valida 5 4 1.43 2.15

Shrimp associations were not only impacted by coral species, but also by coral morphologies. A comparison of preferred corals with and without shrimp occupants indicated that shrimp resided in above average (x̅ = 24.5cm) coral head diameters (Wilcoxon signed rank test for non-normal data, p<0.001, JMP Pro 13) and above average (x̅ = 6.4cm) branch heights (p=0.021), but inter-brachial spacing was not a relevant factor in likelihood of shrimp association (Figure 4.2, n=420). A negative binomial generalized linear model (GLM) was created in R to assess how coral morphology influenced quantity of shrimp within preferred corals. This model was chosen because the data consisted of counts and had an abundance of zeros. Within this model, coral head diameter was the only morphological characteristic that significantly influenced quantity of shrimp within preferred corals (Table 4.2), with shrimp quantity increasing with coral diameter.

Figure 4.2: Habitation of C. graminea within Acropora corals is associated with (A) coral head diameter and (B) coral branch height, but not with (C) inter- branchial spacing. Dark grey bars represent corals without resident C. graminea while light grey bars represent corals with shrimp inhabitants. Wilcoxon signed rank test, a=0.05, * indicates significance.

Table 4.2: Negative binomial generalized linear model to determine influence of coral morphology on quantity of C. graminea.

Source of variation Estimate SE z value p value Diameter 0.058 0.015 3.883 <0.001 Height 0.002 0.098 0.023 0.981 Inter-brachial spacing -0.046 0.486 -0.950 0.342

In addition to shrimp abundance within corals of varying species and morphologies, C. graminea were collected for analyses of fitness and community structure. In all, 44 females, 18 males and 7 juveniles were collected from corymbose or digitate corals. The sex ratio of shrimp within corals was diverse (Figure 4.3), with solitary females (n=6) and solitary males (n=7) occurring frequently. Mixed sexes of one male and one or multiple females and juveniles were found on 10 corals. Multiple

96 females often occupied a single coral colony (n=4); however, multiple males were never found on the same coral. Juveniles whose sex could not be accurately determined were found in isolation or with other males or females. Gravid females were found in corals hosting mixed sexes, solitary females, and multiple females.

Figure 4.3: Sex ratios of C. graminea within Acropora corals. 69 shrimp were collected from 31 coral individuals: 44 females, 18 males, and 7 juveniles.

97 Male C. graminea exhibited smaller SL (t-test in JMP, p=0.015), TL (p=0.019), and carapace width (p=0.047) but comparable claw sizes to females (Figure 4.4). In contrast, the ratio of claw length to width was significantly smaller in males (t- test, p<0.001). Four shrimp (3 females, 1 juvenile) were missing both claws; 5.5% of males, 22.7% of females, and 57.1% of juveniles were missing one claw. Among shrimp that possessed both claws, the size difference between the two claws was much more pronounced in females than in males (t-test, p<0.001). Juveniles were not included in this analysis as only three individuals possessed both claws.

98 Shrimp fitness was assessed through body size (TL, SL, carapace width), defensive strategies (claw length, width, L/W ratio), and fecundity (egg count). Only adult shrimp that could be sexed were used in this analysis to control for life stage (n=62). Additionally, two shrimp hosted the isopod parasite, Neophryxus globicaudatus (Bruce 1973, Bruce 2007), beneath their anterior abdominal segment and were removed from the fecundity analysis, as this parasite may influence egg retention. Body length (SL, TL) was log transformed to normalize the data. A multiple linear regression model was constructed to assess the influence of coral head diameter and branch height on shrimp body characteristics. Since male C. graminea were smaller than females, sex was added as a categorical predictor into the model. Only significant interactive effects were left in the model. Graphical linear regressions are depicted in the supplementary figures listed in Appendix A. Coral diameter, branch height, and sex were all significant predictors of body size (SL, TL and carapace width) (Table 4.3, Figure A.1). The interaction between sex and coral diameter significantly influenced SL and TL, but not carapace width. Inter-brachial spacing was not recorded for 36 corals, so these data were analyzed in a separate regression model, revealing that both inter-brachial spacing and sex were influential factors of shrimp body size (Table 4.4). Additionally, the interaction between inter- brachial spacing and sex was a significant predictor of all three characteristics of body size.

99 Table 4.3: Multiple linear regression results to predict body size characteristics (TL, SL, carapace width). Diameter, height and sex were used as predictors in the model. The only significant interaction effect was between coral diameter and shrimp sex. Regression graphs are depicted in Appendix A. a=0.05, * indicates significance.

Body Size Parameters Predictor Total Length Standard Length Carapace Width

Diameter F1,62=13.92, p<0.001* F1,62=16.68, p<0.001* F1,62=14.75, p<0.001*

Height F1,62=5.94, p=0.018* F1,62=4.08, p=0.048* F1,62=6.78, p=0.012*

Sex F2,62=24.74, p<0.001* F2,62=23.73, p<0.001* F2,62=22.19, p<0.001*

Diameter:Sex F2,62=4.08, p=0.022* F2,62=4.35, p=0.017* F2,62=0.26, p=0.774

Body Size Parameters Predictor Total Length Standard Length Carapace Width Inter-brachial F =7.45, p=0.010* F =6.76, p=0.014* F =4.97, p=0.033* spacing 1,31 1,31 1,31

Sex F1,31=27.51, p<0.001* F1,31=25.87, p<0.001* F1,31=19.27, p<0.001* Inter-brachial F =4.61, p<0.018* F =3.93, p=0.030* F =3.54, p=0.041* spacing:Sex 2,31 2,31 2,31

Influence of coral morphology on claw size (length and width) was also assessed through a multiple linear regression. Coral diameter was an influential factor for mature claw length and width, but coral branch height only impacted claw length (Table 4.5, Figure A2). Sex was a significant factor for both measurements of claw

100 size, and the interaction between branch height and sex was relevant for claw width measurements. Claw L/W ratio was not influenced by coral diameter or height; however, sex and the interaction term were significant predictors. Inter-brachial spacing was not a significant factor in any claw characteristics (Table 4.6).

Claw Size Parameters Predictor Claw Length Claw Width Claw L/W Ratio

Diameter F1,58=18.93, p<0.001* F1,58=14.46, p<0.001* F1,58=0.65, p=0.425

Height F1,58=5.70, p=0.020* F1,58=3.64, p=0.061 F1,58=3.33, p=0.073

Sex F2,58=20.56, p<0.001* F2,58=27.18, p<0.001* F2,58=6.23, p=0.004*

Diameter:Sex F2,58=2.19, p=0.121 F2,58=3.20, p=0.048* F2,58=5.93, p=0.005*

Claw Size Parameters Predictor Claw Length Claw Width Claw L/W Ratio Inter-brachial F =1.77, p=0.193 F =2.04, p=0.163 F =0.001, p=0.971 spacing 1,32 1,32 1,32

Sex F1,32=13.97, p<0.001* F1,32=19.76, p<0.001* F1,32=9.63, p=0.393

101

Fecundity, measured through egg counts of gravid shrimp, was analyzed using a multiple linear regression with height and diameter as predictor variables. The interaction term was not significant and was therefore removed. Coral diameter significantly influenced fecundity (F1,31=5.08, p=0.031), but branch height did not

(F1,31=1.42, p=0.243) (Appendix A). Additionally, inter-brachial spacing was a significant predictor of fecundity; however, the n-value of observations of fecund shrimp within corals with recordings of inter-brachial spacing was low (n=13), thus the power of the test was minimal. Egg quantity was positively correlated with shrimp SL, carapace width, and claw length, but not influenced by TL, claw width or claw L/W ratio (Table 4.7). Finally, shrimp morphometric measurements were correlated with the community of conspecifics within the coral hosts. Carapace width, and claw length and width were positively correlated with the number of resident C. graminea within the coral (Table 4.8) After Bonferroni adjustment, fecundity, SL, TL and claw

L/W ratio were not significantly impacted by quantity of shrimp (a=0.007).

102 Table 4.7: Multiple regression analysis to determine influence of body (TL, SL, carapace width) and claw (length, width, ratio) dimensions on shrimp fecundity measured by egg count. Body size predictors and claw size predictors were analyzed in two separate models. No interaction terms were significant were thus removed from the model. a=0.05, * indicates significance

Fecundity Parameter Predictor Egg count

Total Length F1,30=2.66, p=0.113

Standard Length F1,30=75.91, p<0.001*

Body size Carapace Width F1,30=11.94, p=0.002*

Claw Length F1,28=43.58, p<0.001*

Claw Width F1,28=1.78, p=0.193 Claw size Claw L/W Ratio F1,28=0.60, p=0.444

Body size parameters Claw size parameters Fecundity Standard Total Carapace Claw Claw Claw Predictor Egg count Length Length Width Length Width L/W ratio

# of shrimp F1,60=6.8 F1,60=7.3 F1,60=16.3 F1,57=17.1 F1,57=8.7 F1,57=4.2 F1,33=5.4 inhabitants p=0.012 p=0.009 p<0.001* p<0.001* p=0.005* p=0.044 p=0.027

103 4.5 Discussion Obligate epibionts have evolved to associate within specific habitats that maximize survival. This specificity is not limited only to the coral species but also to intraspecific variations in coral morphological characteristics. The association between shrimp fitness and host morphology can reveal the ecological significance of these associations, especially during changing conditions. Coralliocaris graminea distributions among corymbose or digitate corals was not random and was impacted by both coral species and morphologies. Additionally, shrimp fitness was mainly impacted by coral head size, branch height and number of resident conspecifics. Although the majority of C. graminea were found within A. nasuta, A. millepora, and A. tenuis corals, these associations were not random once coral abundance was incorporated into the expected values. Shrimp abundances were higher than expected within A. humilis, A. nasuta, A. tenuis, and A. valida corals. While C. graminea were found within 7 acroporids in this study, and at least 18 acroporids in other studies, (Miyaki 1968; Bruce 1972,1976, 1977; Patton 1994; Stella et al. 2010; Vytopil and Willis 2011), they may preferentially select for A. humilis, A. nasuta, A. tenuis, and A. valida. This preference may be morphologically driven as each of these corals, except A. tenuis, exhibited above average coral diameters, branch heights, and inter-brachial spacing. Coral colony size was positively correlated with abundance of C. graminea, a pattern that is mirrored in several other invertebrate coral inhabitants (Abele 1976, Abele and Patton 1976, Coles 1980, Enochs 2012). Larger corals host bigger and fitter populations of C. graminea – potentially due to greater access to food. While the precise diet of C. graminea is unknown, they have the appendages required for suspension feeding (Bruce 1976) and have been observed ingesting zooplankton

104 (Patton 1994). A larger coral would increase food availability by allowing more channels through which water, laden with consumable particles and plankton, can flow. Alternatively, the larger coral area would increase the amount of coral tissue and mucus, as well as provide additional habitat for small organisms such as amphipods. Both coral tissue and small invertebrates may provide additional food resources to C. graminea, if suspension feeding is not its sole source of nutrition. The increase in complexity associated with branch height and inter-brachial spacing may provide better protection from predators. Coralliocaris graminea that have been removed from their coral habitat are quickly consumed by reef fish such as Thalassoma sp., and Bruce (1972) postulated that this may be due to their bright coloration. Even within the coral matrix, these shrimp face predation threat and can be targeted by crabs, stomatopods, and small fish (Bruce 1972). However, C. graminea are able to coexist with the other commensal and symbiotic organisms that reside within the branches and incidence of cohabitation is high (pers. obs.). Moreover, an analysis of the gut content of Trapezia crabs, one of the few epibionts large enough to consume C. graminea, reveals that these crabs subsist on a diet of coral mucus, not shrimp (Castro 1976). Despite the protection that the coral matrix offers, C. graminea still face predation. The frequency of missing or partially regenerated claws is high, which is likely due to predation attempts (Bruce 1972) but could also be a result of competition or mating. Unlike most true snapping shrimp, which have one large claw (Johnson 1947), both claws of C. graminea are modified to produce high-velocity “punches” for protection. These shrimp also use both claws to grasp the coral branches such that they are oriented facedown inside the coral (Patton 1966). Large claw size increased with

105 coral diameter and branch height, suggesting that high rates of resource acquisition allowed for the investment in robust defensive structures. Coralliocaris gramminea have evolved to have two large claws, therefore size discrepancies between the large and small claws likely reflect recent limb loss and subsequent regeneration. This study determined that females not only displayed a larger size discrepancy between claw sizes, but also showed higher rates of single or double limb loss, potentially indicative of either slower rates of recovery or more frequent limb loss in females. Whether limb loss is a function of predation or competition remains to be determined. Females were more common than males, and the sex ratio within individual coral colonies varied from solitary males and females to mixed sexes of one male and multiple females. Gravid shrimp were not only located in corals with both males and females, but also in corals hosting solitary or multiple females only. Movement of males between colonies may be occurring, although the life history and reproductive ecology of these cryptic shrimp remain unknown. As corals continue to decline and die, these obligate shrimp may be forced to relocate to healthy hosts or even non-host species, although movement of C. graminea between corals has not yet been documented. In an investigation of the frequency of obligate coral hosts associating with dead corals, Head et al. (2015) observed only one

C. graminea residing within a dead Acropora. The current study indicates that these specialized shrimp can inhabit several species of structurally-similar Acropora corals, yet specific coral morphologies can significantly influence shrimp abundance and fitness. Therefore, in a system barren of acroporids, these obligate coral associates may be able to survive on non-host corals, provided that the alternate hosts possess similar corymbose or digitate morphologies. Most significantly, shrimp abundance and

106 fitness increased with coral head diameter, highlighting the need for conservation efforts aimed at increasing individual colony size along with overall coral coverage.

4.6 Acknowledgements I would like to thank P. Leingang and A. Good for their assistance with data collection and D Miller for statistical support. I would also like to thank the members of the Dixson Lab for editorial comments as well as the staff of Coralview Island Resort and Matava Resort for logistical support.

107 REFERENCES

Abele LG. 1976. Comparative species richness in fluctuating and constant environments: coral-associated decapod crustaceans. Science. 192:461-463.

Abele LG, Patton WK. 1976. The size of coral heads and the community biology of associated decapod crustaceans. J Biogeogr. 3:35-47.

Anthony KR, Maynard JA, Diaz-Pulido, Mumby PJ, Marshall PA, Cao L, Hoegh- Guldberg OV. 2011. Ocean acidification and warming will lower coral reef resilience. Glob Change Biol. 17:1798-808.

Bruce AJ. 1972. A review of information upon the coral hosts of commensal shrimps of the subfamily Pontoniinae, Kingsley, 1878 (Crustacea, Decapoda, Palaemonidae). J Mar Biol Ass India. 1:339-418.

Bruce A. 1973. Prophryxus globicaudatus gen.nov., sp.nov., a hemiarthrinid bopyrid parasite of pontoniinid shrimps of the genus Coralliocaris Stimpson. Parasitology. 67:365-373.

Bruce AJ. 1976. A report on a small collection of shrimps from the Kenya National Marine Parks at Malindi, with notes on selected species. Zool Verh. 145:1-72.

Bruce AJ. 1977. The hosts of the coral-associated Indo-West-Pacific pontoniine shrimps. Atoll Res Bull. 205:1-19.

Bruce AJ. 1998. New keys for the identification of Indo-West Pacific coral associated pontoniine shrimps, with observations on their ecology (Crustacea: Decapoda: Palaemonidae). Ophelia. 49:29-46.

Bruce AJ. 2007. Neophryxus, a new name for prophryxus bruce, 1973 (Crustacea: Isopoda: Bopyridae). Zootaxa. 1646:67-68.

Bruno JF, Selig ER. 2007. Regional decline of coral cover in the Indo-Pacific: timing, extent, and subregional comparisons. PLoS one. 2:e711.

Castro P. 1976. Brachyuran Crabs Symbiotic with Scleractinian Corals: A Review of their Biology. Micronesica. 12:99-110.

108 Coles SL. 1980. Species diversity of decapods associated with living and dead reef coral Pocillopora meandrina. Mar Ecol Prog Ser. 2:281-291.

Comeau S, Edmunds PJ, Spindel NB, Carpenter RC. 2014. Fast coral reef calcifiers are more sensitive to ocean acidification in short‐term laboratory incubations. Limnol Oceanogr. 59:1081-1091.

Cronin G. 2001. Resource allocation in seaweeds and marine invertebrates: chemical defense patterns in relation to defense theories. In: Marine Chemical Ecology. McClintock JB, Baker BJ (Eds.) CRC Press, Boca Raton, FL. pp.325-353.

Enochs IC. 2012. Motile cryptofauna associated with live and dead coral substrates: implications for coral mortality and framework erosion. Mar Biol. 159:709- 722.

Gardner TA, Côté IM, Gill JA, Grant A, Watkinson AR. 2003. Long-term region-wide declines in Caribbean corals. Science. 301:958-960.

Head CE, Bonsall MB, Koldewey H, Pratchett MS, Speight M, Rogers AD. 2015. High prevalence of obligate coral-dwelling decapods on dead corals in the Chagos Archipelago, central Indian Ocean. Coral Reefs. 34:905-915.

Hughes TP, Tanner JE. 2000. Recruitment failure, life histories, and long‐term decline of Caribbean corals. Ecology. 81:2250-2263

Hughes TP, Rodrigues MJ, Bellwood DR, Ceccarelli D, Hoegh-Guldberg O, McCook L, Moltschaniwskyj N, Pratchett MS, Steneck RS, Willis B. 2007. Phase shifts, herbivory, and the resilience of coral reefs to climate change. Curr Biol. 17:360-365.

Hughes TP, Kerry JT, Baird AH, Connolly SR, Dietzel A, Eakin CM, Heron SF, Hoey AS, Hoogenboom MO, Liu G, McWilliam MJ. Global warming transforms coral reef assemblages. Nature. 556:492-496.

Javahery S, Nekoubin H, Moradlu AH. 2012. Effect of anaesthesia with clove oil in fish. Fish Physiol Biochem. 38:1545-1552.

JMP Pro, Version 13, SAS Institute Inc., Cary, NC, 1989-2007.

Johnson MW, Everest FA, Young RW. 1947. The role of snapping shrimp (Crangon and Synalpheus) in the production of underwater noise in the sea. Biol Bull. 93:122-138.

109 Jones GP, McCormick MI, Srinivasan M, Eagle JV. 2004. Coral decline threatens fish biodiversity in marine reserves. Proc Natl Acad Sci USA. 101:8251-8253.

Loya Y, Sakai K, Yamazato K, Nakano Y, Sambali H, van Woesik R. 2001. Coral bleaching: the winners and the losers. Ecol Lett. 4:122-131.

MacArthur R, Levins R. 1964. Competition, habitat selection, and character displacement in a patchy environment. Proc Natl Acad Sci USA. 51:1207- 1210.

Miyake S, Fujino T. 1968. Pontoniid shrimps from the Palau Islands (Crustacea, Decapoda, Palaemonidae). J Fac Agr Kyushu U. 14:399-431.

Munday PL. 2004. Habitat loss, resource specialization, and extinction on coral reefs. Glob Change Biol. 10:1642-1647.

Pandolfi JM, Connolly SR, Marshall DJ, Cohen AL. 2011. Projecting coral reef futures under global warming and ocean acidification. Science. 333:418-422.

Patton WK. 1966. Decapod Crustacea commensal with Queensland branching corals. Crustaceana. 10:271-295.

Patton WK. 1994. Distribution and ecology of animals associated with branching corals (Acropora spp.) from the Great Barrier Reef, Australia. Bull Mar Sci. 55:193-211.

R Core Team. 2017. R: A language and environment for statistical computing. R Foundation for Statistical Computing, Vienna, Austria. URL: https://www.R- project.org/.

Schindelin, J, Arganda-Carreras I, Frise E, Kaynig V, Longair M, Pietzsch T, Preibisch S, Rueden C, Saalfeld S, Schmid B, Tinevez JY. 2012. Fiji: an open- source platform for biological-image analysis. Nature Methods 9:676-682, PMID 22743772, doi:10.1038/nmeth.2019

Stella JS, Jones GP, Pratchett MS. 2010. Variation in the structure of epifaunal invertebrate assemblages among coral hosts. Coral Reefs. 29:957-973.

Stella JS, Pratchett MS, Hutchings PA, Jones GP. 2011. Coral-associated invertebrates: diversity, ecological importance and vulnerability to disturbance. Oceanogr Mar Biol Ann Rev. 49:43–104

Vytopil E, Willis B. 2001. Epifaunal community structure in Acropora spp.(Scleractinia) on the Great Barrier Reef: implications of coral morphology and habitat complexity. Coral Reefs. 20:281-288.

110 Wild C. Hoegh-Guldberg O, Naumann MS, Colombo-Pallotta MF, Ateweberhan M, Fitt WK, Iglesias-Prieto R, Palmer C, Bythell JC, Ortiz JC, Loya, Y. 2011. Climate change impedes scleractinian corals as primary reef ecosystem engineers. Mar Freshwater Res. 62:205-215.

Williams DE, Miller MW. 2012. Attributing mortality among drivers of population decline in Acropora palmata in the Florida Keys (USA). Coral Reefs. 31:369- 382.

111 Chapter 5

CONCLUDING REMARKS

Through an analysis of coral larvae and habitat-specialized shrimp, I have investigated critical aspects of habitat selection in coral reef invertebrates. Habitat selection is vital at many points throughout an organism’s life and can be mediated by numerous environmental cues, depending on the sensory capabilities and ecological requirements of the species. Regardless of timing or scope, selection of habitat can have pervasive impacts on organismal fitness. Caribbean scleractinian coral larvae are invariably attracted to general reef cues throughout their ontogeny but reveal diverse behaviors regardless of cue. Observations of minute behaviors revealed that directed movement and small circles might be indicative of positive benthic cues. Appropriate habitat selection is essential for coral larvae because corals only have a single opportunity to make the optimal decision before becoming permanently sessile after metamorphosis (Hadfield 1986). The significance of this decision is accentuated by the unmitigated importance of corals as a foundational species in the reef ecosystem (Graham and Nash 2013).

Successful recruitment of coral larvae is vital to reef recovery (Dixson et al. 2014). Habitat selection is not limited to the larval stage of reef invertebrates but can also be relevant for homing adults. Peppermint shrimp, Lysmata pederseni, rely on both chemical and morphological cues to select appropriate tube sponges after nightly forays on the reef. Invertebrates that rely on other living organisms for habitat are at a heightened vulnerability, especially during periods of environmental change. Not only

112 can specialists be directly threatened by change, but they can also be indirectly affected if their host cannot tolerate the stress (Wilson et al. 2008). The appropriateness and health of a host can impact the fitness of inhabitants, as investigated in the shrimp, Coralliocaris graminea. As morphology is potentially more relevant than habitat species in maintaining fitness, the potential exists for these habitat specialists to adapt to environmental change by transferring to hardier, yet morphological similar, non-host corals. Coral reefs are declining at an unprecedented rate due to local and global stressors (Knowlton and Jackson 2008). While the potential remains for local stressors to be mitigated by community efforts (Cinner et al. 2005; Hamilton et al. 2011), global stressors, such as climate change and ocean acidification, will continue to exacerbate the deterioration (Hoegh-Guldberg et al. 2007). Projected reductions in ocean pH can have devastating impacts on the sensory biology of marine organisms, sometimes completely reversing ecological behaviors critical for survival (Ashur et al. 2017; Appendix B). All of the behaviors discussed in the previous chapters have the potential to be influenced by this change in ocean chemistry. Continuing to explore the influences of environmental stimuli on habitat selection in reef invertebrates is an important foundational step in coral reef ecology. From there, we can advance our knowledge of how these behaviors might be affected by changing conditions so that we can aim to effectively protect the vital coral reef ecosystem.

113 REFERENCES

Ashur MM, Johnston NK, Dixson DL. 2017. Impacts of ocean acidification on sensory function in marine organisms. Integrative and comparative biology. 2017. 57:63-80.

Cinner JE, Marnane MJ, McClanahan TR. 2005. Conservation and community benefits from traditional coral reef management at Ahus Island, Papua New Guinea. Conserv Biol. 19:1714-1723.

Dixson DL, Abrego D, Hay ME. 2014. Chemically mediated behavior of recruiting corals and fishes: a tipping point that may limit reef recovery. Science. 345:892-897.

Graham NA, Nash KL. 2013. The importance of structural complexity in coral reef ecosystems. Coral Reefs. 32:315-26.

Hadfield MG. 1986. Settlement and recruitment of marine invertebrates: a perspective and some proposals. Bull Mar Sci. 39:418-25.

Hamilton RJ, Potuku T, Montambault JR. 2011. Community-based conservation results in the recovery of reef fish spawning aggregations in the Coral Triangle. Biol Conserv. 144:1850-1858.

Knowlton N, Jackson JB. 2008. Shifting baselines, local impacts, and global change on coral reefs. PLoS Biology. 6:e54.

114 Appendix A

SUPPLEMENTARY FIGURES

2.5

1.5

Total Length (cm) 0.5

2.5

1.5

0.5 Standard Length (cm) 0 0.8

0.6

0.4

0.2 Carapace Width (cm) 0 0 20 40 60 80 0 2 4 6 8 10 12 Coral Head Diameter (cm) Coral Branch Height (cm) Females Males Figure A.1: Linear regression graphs displaying relationship between body size characteristics (TL, SL, carapace width; y-axis) and coral morphology (diameter, height; x-axis). Female shrimp are depicted with red markers while males are depicted with blue markers. Full statistical outputs are described in Figure 4.3.

115

1.5

1 Claw Length (cm) 0.5

0.5

0.4

0.3

0.2 Claw Width (cm) 0.1

0 0 20 40 60 80 0 5 10 15 Coral Head Diameter (cm) Coral Branch Height (cm) Females Males Figure A.2: Linear regression graphs displaying relationship between claw size characteristics (length, width, y-axis) and coral morphology (diameter, height; x-axis). Female shrimp are depicted with red markers while males are depicted with blue markers. Full statistical outputs are described in Figure 4.5.

116 600 500 400 300 200

Number of eggs 100 0 0 20 40 60 80 0 2 4 6 8 10 12 Coral Head Diameter (cm) Coral Branch Height (cm)

Figure A.3: Linear regression graphs displaying relationship between fecundity (number of eggs) and coral morphology (diameter, height; x-axis). Full statistical outputs are described in Figure 4.5.

117 Appendix B

IMPACTS OF OCEAN ACIDIFICATION ON SENSORY PERCEPTION: UNDERLYING MECHANISMS AND TAXA COMPARISONS

This appendix has been published in the journal Integrative and Comparative Biology. Co-Authors: Nicole K. Johnston1, Danielle L. Dixson1,2 Addresses: 1School of Biology, Georgia Institute of Technology, 310 Ferst Drive, Atlanta, GA, 30363, USA 2School of Marine Science and Policy, University of Delaware, 700 Pilottown Road, Lewes, DE, 19958, USA

B.1 Introduction The global decline of oceanic ecosystems has been attributed to various natural and anthropogenic stressors, with ocean acidification identified as a major threat to marine life. Ocean acidification is caused by the influx of atmospheric CO2 into the oceans, resulting in an increase in pCO2 and reduction of surface water pH (Doney et al. 2009). This change in acidity has significant impacts on the physiology (Fabry et al. 2008), ontogeny (Byrne 2011), behavior (Clements and Hunt 2015; Nagelkerken and Munday 2016) and survivorship (Kroeker et al. 2010) of marine organisms, which can escalate to the ecosystem level. Recently, numerous studies covering a myriad of taxa have observed an impairment of organismal sensory perception when exposed to ocean acidification conditions (Munday et al. 2009; Dixson et al. 2010).

118 Sensory stimuli allow organisms to perceive their external environment. Perception of chemical, auditory, and visual cues plays an integral role in the daily life and survival of marine organisms by influencing homing (Simpson et al. 2005; Dixson et al 2008), settlement (Pawlik et al. 1992), predator detection and evasion (Lima and Dill 1990), foraging (Tomba et al. 2001), conspecific social interactions (Cole and Shapiro 1995) and mating (Thomas 2011). Additional senses - including the lateral line system, electrosense in elasmobranchs and the ability to detect magnetic fields - offer unique conduits of perceiving the world to specialized marine species. A critical distinction exists when describing sensory ecology that becomes a challenge when designing experiments – sensory input versus central processing. Sensory input is the process of transmitting impulses from sensory organs to nerve centers at which point central processing converts the stimuli into behavior (The American Heritage Medical Dictionary 2007). Studies that strictly examine sensory input accurately reveal the proximate causes of the affected behaviors. However, the majority of studies assessing sensory perception under ocean acidification use observable behaviors as a proxy for sensory perception, yet often fail to distinguish whether the behavioral differences are a result of distorted sensory input or altered central processing, or both. In this study, we attempt to characterize relevant studies based on the true message each experiment reveals. At the organismal scale, ocean acidification directly affects the perception of a suite of sensory systems, including chemosensory (Munday et al 2009), vision (Ferrari et al. 2012a) and sound (Simpson et al. 2011) and additional daily cognitive functions including activity level and boldness (Munday et al. 2010), lateralization (Domenici et al. 2011) and learning (Ferrari et al. 2012b). These disturbances have the potential to

119 reverberate through the ecosystem by influencing recruitment (Ferrari et al. 2011), settlement (Devine et al. 2012a), replenishment (Munday et al. 2010), connectivity (Munday et al. 2009) and predator-prey interactions (Manriquez et al 2014). This review aims to summarize and analyze the current knowledge on the effect of ocean acidification on sensory systems in marine organisms. We explore chemosensory perception, vision, and auditory detection in the context of (1) direct sensory studies, (2) possible mechanisms and (3) expected differences between taxa. We also address other sensory systems including electroreception, magnetic sense and the lateral line system and hypothesize why elevated CO2 may also influence these systems and advocate for additional studies. We aim to not only summarize the breadth of current research but also identify biases, discrepancies, patterns, gaps in the knowledge, and areas of future study.

B.2 Chemosensory Perception

The ability to detect chemical cues is the primary sense used by most organisms across different environments and taxa (Ache and Young 2005). Recognition of chemical cues is initiated by the binding of a chemical molecule to a receptor protein on a primary chemoreceptor neuron. In all animals, this binding initiates a G protein-based signaling cascade that transmits a signal to the central nervous system (Buck and Axel 1991). While the physiology of chemosensory perception is relatively conserved across species, the morphology of chemoreception organs can vary greatly; crustaceans use aesthetascs on their antennules to “sniff” their environment (Schmitt et al. 1979) and fish and cephalopods use specialized olfactory organs exposed to chemical cues through nasal cavities or olfactory pits (Kasumyan

120 2004; Polese et al. 2016). It is still debated whether the reception of chemical signals is through gustation or olfaction (Hara 1994). Therefore in the context of this review, the term chemosensory perception is used when referring to either sense. Chemosensory perception and communication are critical in the marine environment and are relevant at all spatial scales (Hay 2009). The ability to accurately detect and respond to environmental chemical cues is critical for homing (Atema et al. 2002; Dixson et al. 2014), settlement (Lecchini et al. 2005), foraging (Tomba et al. 2001) and predator evasion (Lima and Dill 1990). The highly conserved nature of chemosensory perception has led to numerous studies investigating the effect of ocean acidification on this essential sense across many taxa. Chemosensory perception and associated behavioral responses are influenced by elevated CO2 (Table A.1), although the physiological or biochemical origin of these changes varies among taxa.

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Sensitivity Study Species CO2 (pCO2) Control (pCO2) CO2 Exposure (pCO2) Munday et al. 2009 Amphiprion percula 1000, 1700 ppm 380 ppm 1000 ppm 11d; settlement* Albright et al. 2010 Acropora palmata 560, 800 μatm 400 uatm 560 μatm 6d; settlement 24h, NH, 11d, Dixson et al. 2010 Amphiprion percula 1000 ppm 380 ppm 1000 ppm settlement*

Munday et al. 2010 Amphiprion percula, Pomacentrus wardi 550, 700, 850 ppm 390 ppm 700 ppm 1-10d*; 1-4d; settlement

122 Albright and Langdon 2011 Porites astreoides 560, 800 μatm 380 μatm 560 μatm 2d; settlement* Cripps et al. 2011 Pseudocromis fuscus 600, 950 μatm 400 μatm 600 μatm 4-7d; adult

Pomacentrus chrysurus, P. moluccensis, Ferrari et al. 2011 700, 850 ppm 390 ppm 700 ppm 4d; settlement P. amboiensis, P. nagasakiensis de la Haye et al. 2012 Pagurus bernhardus pH 6.8 pH 8.20 pH 6.8 5d; adult Pomacentrus amboinensis, P. chrysurus, Devine et al. 2012a 700, 850 ppm 440 ppm 700 ppm 4-5d; settlement P. moluccensis Devine et al. 2012b Cheilodipterus quinquelineatus 550, 700, 950 ppm 390 ppm 550 ppm 4d; adult Doropoulos et al. 2012 Acropora millepora 800, 1300 μatm 401 μatm 800 μatm 6d; settlement Amphiprion percula, Neopomacentrus Nilsson et al. 2012 900 μatm 450 μatm 900 μatm 11d; 18d; settlement azysron

Chivers et al. 2013 Pomacentrus amboiensis 987 μatm 440 μatm 987 μatm 4d; juvenile

Devine and Munday 2013 Paragobiodon xanthosomus 880 μatm 440 μatm 880 μatm 4d; adult

Jutfelt and Hedgarde 2013 Gadus morhua 1170 μatm 550 μatm No effect 35-41d; juvenile

Lönnstedt et al. 2013 Pomacentrus amboiensis 880 μatm 440 μatm 880 μatm 4d; settlement

Munday et al. 2013 Plectropomus leopardus 570, 700, 960 μatm 490 μatm 700 μatm 4d, 28 d; juvenile CCA: 6 weeks; Webster et al. 2013 Coral larvae/CCA 822, 1187, 1638 μatm 464 μatm 822 μatm settlement Barry et al. 2014 Strongylocentrotus fragilis 3255 ppm 1028 ppm 3255 ppm 30d; adult Dixson et al. 2014 Mustelus canis 741, 1064 μatm 405 μatm 1064 μatm 5d; adult Manriquez et al. 2014 Concholepas concholepas 700, 1000 μatm 390 μatm 1000 μatm 5mo; juvenile Dascyllus aruanus, Pomacentrus Munday et al. 2014 moluccensis, Apogon cyanosoma, 441-998 μatm 346-413 μatm No effect Entire life; juvenile* Cheilodipterus quinquelineatus Welch et al. 2014 Acanthochromis polyacanthus 656, 912 μatm 446 μatm 656 μatm 40-45d; juvenile* Queiros et al. 2015 Nucella lapillus 750, 1000 μatm 380 μatm 1000 μatm 14mo; adult 1 23

Many marine organisms have a dispersive larval stage followed by a relatively sessile adult stage (Caley et al. 1996). Most development occurs in the pelagic environment and the successful return and recruitment of juveniles is vital for species persistence in any marine ecosystem. When in the pelagic environment, larvae primarily utilize chemoreception (Atema et al. 2002) and sound (Simpson et al. 2005) to navigate towards a suitable habitat. For example larval anemonefish, Amphiprion percula, use the chemical signatures of island leaf litter to navigate to reef systems supporting their symbiotic host anemones (Dixson et al. 2008). After exposure to future ocean acidification conditions, however, A. percula displayed a weaker positive reaction to native trees, indicative of reef habitat, and significant positive response to non-native plants, a cue not correlated to beneficial habitat (Munday et al. 2009). Additionally, larval damselfish (family Pomacentridae) in ambient conditions have specific habitat cue preferences, depending on the species. However, once exposed to lower pH, each species was unable to distinguish between odor cues of different habitat when the chemical cues alone were tested, however, settlement was successful when all cues were present, suggesting sensory compensation (Devine et al. 2012a). Accurate settlement decisions are especially critical for sessile invertebrates, as habitat choice is permanent once settlement and metamorphosis have occurred.

Invertebrate larvae use chemoreception to choose specific habitat locations and settlement can be induced by an extensive array of cues allowing gregarious (i.e. conspecific cues) and associative (i.e. parasitic, herbivorous, microbial cues) settlement (Pawlick 1992). Ocean acidification reduces the percentage of successful settlement of several species of coral, including Acropora millepora (Webster et al. 2013; Doropoulos et al. 2012), A. tenuis (Webster et al. 2013), A. palmata (Albright et

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al. 2010), and Porites astreoides (Albright and Langdon 2011). Additionally, under elevated CO2 conditions, A. millepora switches preferred substrate preference (Doropoulos et al. 2012). Thus far, no study has tested the effects of ocean acidification on chemoreception in isolation, i.e. within a chemical choice flume so that only chemical cues are assessed; it is therefore unknown if the reduction in settlement success is a function of chemosensory impairment. The reduction in settlement may be related to a change in microbial composition on preferred settlement substrates. The microbial shift may change the chemical signature, resulting in reduced settlement success (Webster et al. 2013). Habitat selection is also important for other life stages beyond recruits. Chemosensory mediated impairment of habitat selection in adult fish is evidenced by

(1) adult gobies, Paragobiodon xanthosomus, that after high CO2 exposure no longer show strong preference for their sole coral host and neglect to relocate to preferred habitat if placed on a dead coral (Devine and Munday 2013) and (2) CO2-exposed adult cardinalfish, Cheilodipterus quinquelineatus, demonstrate impairment in the ability to navigate back to their resting sites after nocturnal feeding (Devine et al. 2012b). The effect of ocean acidification on the navigational capabilities of adult invertebrates has not been assessed.

Beyond homing and settlement, chemosensory cues are vital for predator detection and evasion. Innate predator recognition is imperative for larvae that recruit to predator-rich habitats such as reefs. Studies on several species of juvenile reef fish (Families: Pomacentridae, Apogoninae, Serranidae) show that under ocean acidification conditions, recognition of predator odor is reversed or the antipredator response behavior is reduced (Dixson et al. 2010; Munday et al. 2010; Munday et al.

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2013; Nilsson et al. 2012; Munday et al. 2014; Welch et al. 2014; Lonnstedt et al. 2013; Chivers et al. 2014; Ferrari et al. 2012a; Ferrari et al. 2011). Larval fishes preferentially swim towards a predator or no longer show preference away from the threat. In contrast, anemonefish, A. percula, when exposed to elevated CO2 conditions only during the egg stage maintained predator avoidance behavior post hatching, suggesting adaptation potential relating to natural variations of pH and CO2 within the reef matrix or the higher levels of CO2 within the egg itself due to embryonic respiration (Dixson et al. 2010). Altered chemosensory perception of predators is not observed in all species of fishes, with juvenile Atlantic cod (Gadus morhua) avoiding predator cues even after month-long high CO2 exposure (Jutfelt and Hedgarde 2013). As for invertebrates, predator avoidance behaviors in two species of Mollusca, including the fast-escape jump response of Gibbosus gibbosus (Watson et al. 2014) and the ability of Concholepas concholepas to distinguish a predator odor from a neutral cue (Manriquez et al. 2014), were impaired by ocean acidification. The physical escape responses of these gastropods were not affected, i.e. G. gibbosus could still jump, however the chemosensory or decision-making capabilities of the organisms were impaired. Predators are also affected by ocean acidification. Chemosensory tracking or attraction to prey cues is diminished after exposure to elevated CO2 in an adult elasmobranch (smooth dogfish, Mustelus canis; Dixson et al. 2015) and a teleost (brown dottyback, Pseudochromis fuscus; Cripps et al. 2011). Foraging ability in adult invertebrates appears to be weakened, by elevated CO2 in species of Mollusca (Queirós et al. 2015), Echinodermata (Barry et al. 2014) and Crustacea (de la Haye et al. 2012), although the specific parameters tested are variable among research groups.

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In the deep ocean, Barry et al. (2014) documented that deep-sea urchins,

Strongulocentrotus fragilis, exposed to elevated CO2 experienced significantly longer foraging time, but not a change in speed, indicating difficulty locating the cue rather than locomotor impairment. Conversely, the gastropod, Nucella lapillus, did not experience increased foraging time, but the distance covered was significantly greater

(Queirós et al. 2015). The hermit crab, Pagurus bernhardus, spent significantly less time in contact with the prey cue and experienced reduced anntenular flicking (“sniffing”) rates (de la Haye et al. 2012). In contrast, juvenile muricid snails, Concholepas concholepas, show no impaired ability to locate prey (Manriquez et al. 2014). Each of these experiments incorporated chemoreception and implied impairment, but chemosensory cues in isolation have yet to be tested. Some studies have implied that in future predator-prey interactions, both players will be affected by reduced pH, but ocean acidification will have a more deleterious impact on prey than predators. Manriquez et al. (2104) tested C. concholepas as both a predator and a prey and found that predator avoidance behavior was reduced by ocean acidification, but prey detection was unaffected. Cripps et al. (2011) found that a mesopredator spent 20% less time in prey cue, which is disadvantageous, but minimal compared to the 80% (check) switch observed in prey species (Dixson et al. 2010). Other factors outside of sensory impairment and behavioral alterations can influence future predator-prey interactions, including growth, shell thickness and strength, strength of offensive weapons and reproduction (Kroeker et al. 2014). After taking into account the effects on both predators and prey, Kroeker et al. (2014) predicts that per capita predation rates of coastal molluscs will increase with ocean acidification. Evolutionarily, it is far more detrimental to have

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reduced predator avoidance than prey detection because the worst result is death rather than undernourishment. Therefore, we can hypothesize that selection pressure will be higher for prey species, possibly suggesting evolutionary adaptation. Studies comparing sensory impairment from both the predator and prey perspective are necessary to scale-up the effects of ocean acidification to the ecosystem level.

B.3 Audition

Audition in the marine environment depends on the definition of sound detection (Budelman 1992). If broadly defined as the ability to detect non-contact vibrational stimuli (Pumphrey 1950), then response to auditory cues has been observed in fish (Popper and Fay 1973), arthropods (Lovell 2005; Derby 1982), cephlopods (Kaifu 2008) and cnidaria (Vermeij 2010). Auditory morphology in fish is similar to that observed in other vertebrates, with the majority of sound detection dependent on the inner ear, which consists of three chambers containing sensitive hair cells (Popper and Fay 1973). When sound waves pass through a fish, the otoliths move within their fluid chambers and press against the hairs, sending impulses to the brain. Marine invertebrates lack inner ears, but can sense vibrational stimuli through other structures, including external sensory hairs in lobsters (Derby 1982), statocysts (gravistatic organs) in shrimp (Lovell et al. 2005) and squid (Kaifu 2008) or external cilia in coral larvae (Vermeij et al. 2010). Sound production and detection is ecologically important for conspecific communication, foraging, predator detection, and navigation (Myrberg 1997). Ecosystem-specific sounds, both biologic, i.e. fish vocalization or crackle of snapping shrimp (Simpson et al. 2010), and physical, i.e. wave action (Urick 1984), can be

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detected from thousands of meters away and can be a reliable source of information for homing fish as sound transmits through water regardless of water movement (Simpson et al. 2005). Auditory cues from the marine environment influence the settlement of crab zoea and megalopae (Jeffs et al. 2003), temperate blennies (Tolimieri et al. 2000), coral larvae (Vermeij et al. 2010) and coral reef fish (Leis et al. 2002, Simpson et al. 2005). Ocean acidification alters this ability of fish larvae to navigate using acoustic cues, affecting both the timing of settlement (Simpson et al. 2011) and the attraction to preferred habitat cues (Rossi et al. 2015, 2016a) (Table A2). In ambient conditions, settlement-stage fish are attracted to nocturnal reef sounds but repelled by daytime, predator-rich reef sounds (Heenan 2012, Leis 1996). However, when exposed to elevated CO2, larval anemonefish, A. percula, experience an attraction to daytime reef noises, which could have implications for survival (Simpson et al. 2011). Similar tests were conducted on temperate mulloway (Argyrosomus japonicas) (Rossi et al. 2016a) and the catadromous fish, barramundi (Lates calcarifer) (Rossi et al. 2015). In present- day conditions, these fishes are attracted to acoustic habitat cues, but avoid them once treated with lower pH. Not only are auditory capabilities altered in mulloway exposed to ocean acidification, but also the soundscape of the preferred habitat is degraded as a result of a reduced intensity and frequency of snapping shrimp sounds (Rossi et al. 2016b). Therefore, ocean acidification has the potential to directly, through sensory impairment, and indirectly, through soundscape depreciation, affect navigation of settlement-stage fish.

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Sensitivity Study Species CO2 (pCO2) Control (pCO2) CO2 Exposure (pCO2) Auditory perception 17-20d; post- Simpson et al. 2011 Amphiprion percula 600, 700, 900 μatm 390 μatm 600 μatm settlement* Otolith development Checkley et al. 2009 Atractoscion nobilis 993, 2558 μatm 380 μatm 933 μatm 7-8d; larvae*

1 1260, 1859, 2626, 30 Franke and Clemmensen 2011 Clupea harengus L. 480 μatm No effect NH* 2903, 4635μatm

Munday et al. 2011a Amphiprion percula 1050, 1721 μatm 404 μatm 1721 μatm 11d; settlement* Munday et al. 2011b Acanthochromis polyacanthus 600, 725, 850 μatm 450 μatm No effect 21d; juvenile* 800, 2100, 3500, 5400 Bignami et al. 2013a Rachycentron canadum 300 μatm 800 μatm 20d; larvae μatm 800, 2100, 3500, 5400 Bignami et al. 2013b Rachycentron canadum 300, 500 μatm 800 μatm 14,22d; larvae μatm Maneja et al. 2013 Gadus morhua 1800, 4200 μatm 370 μatm 1800 μatm 7-53d; larvae* Schade et al. 2014 Gasterosteus aculeatus 1000 μatm 400 μatm 1000 μatm 30,60,90d; larvae* Perry et al. 2015 Stenotomus chrysops 1726; 1861 μatm 1205 μatm No effect 8w; adult Reveillac et al. 2015 Sparus aurata 700, 1200, 2000 μatm 475 μatm 700 μatm 40d; juvenile

Impairments to the auditory system in fish suggest that sensory systems with internal receptors are affected by changing ocean chemistry. Whereas olfactory receptors in most organisms are located externally, fish auditory capabilities are dependent on internal structures such as the inner ear and otoliths. A change in shape, size or symmetry of otoliths can be detrimental to organisms (Gagliano et al. 2008).

While elevated-CO2 induced alterations in otolith symmetry have not been observed (Munday et al. 2011a,b; Maneja et al. 2013; Reveillac et al. 2015), an increase in otolith size in fish exposed to elevated CO2 conditions has been recorded in numerous species (Checkley et al. 2009, Schade et al. 2014, Bignami et al. 2013a,b, Munday et al. 2011a, Hurst et al. 2012, Maneja et al. 2013, Frommel et al. 2013, Reveillac et al. 2015, Rossi et al. 2016) (Table A.2), suggesting a possible mechanism for auditory alteration. In contrast, the otoliths of some species remain unaffected by even high levels of CO2 (Munday et al. 2011b, Franke and Clemmesen 2011, Simpson et al. 2011, Perry et al. 2015), indicating species-specific sensitivity. An increase in otolith size corresponds to more sensitive hearing and models predict up to a 50% increase in hearing range for CO2-treated fish larvae due to enlarged otoliths (Bignami et al. 2013a). An improvement in audio detection could be beneficial for perceiving ecologically important cues, however, could be disadvantageous through increased sensitivity to background noise, therby obscuring relevant audio information (Bignami et al. 2013a). Further, the benefits or detriments of increased sensitivity could be species dependent. Evolution of benthic organisms has already selected for comparatively large otoliths, thus these species would likely benefit from increased sensitivity (Lychakov and Rebane 2000). However, pelagic fish developed small otoliths and an incrase in sound detection may distract or confuse these less sensitive

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organisms (Lychakov and Rebane 2000). Typically, the level of CO2 exposure needed to increase otolith size (avg levels) is higher than levels needed to induce altered auditory perception (avg levels). Therefore, behavioral effects will most likely occur sooner than otolith enlargement, lessening the likelihood that otolith enlargement is responsible for altered perception.

B.4 Vision

Vision in marine organisms is remarkably diverse, with visual structures ranging from pigmented photoreceptors in some invertebrates and early chordates to the complex and highly functional camera eyes found in fish and cephalopods (Arendt 2003). Morphologically, there are dozens of different eye types, with almost every form existing in the aquatic environment (Land and Nilsson 2002). In aquatic systems, the ability to focus an image depends on the formation of either pinhole eyes or lenses (Land and Fernald 1992). The physiological basis of vision relies on the photon- activated G-protein phototransduction cascade that is present in nearly every light- sensing organism (Arshavsky et al. 2002). Despite variations among taxa, this cascade is overall conserved. While auditory and olfactory cues are integral for navigation across large spatial scales, vision becomes critical over a smaller geographic area once an organism has located a specific habitat. Depending on conditions, visual cues can provide the most accurate information about the environment, particularly predation risk (Ferrari et al. 2010). Several studies have indicated deficient response to visual cues and altered retinal functioning under ocean acidification conditions (Table A.3).

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Sensitivity Study Species CO2 (pCO2) Control (pCO2) CO2 Exposure Visual components (pCO2) Ferrari et al. 2012a Pomacentrus amboiensis 550, 700, 850 μatm 440 μatm 850 μatm 4d; settlement Visual perception Ferrari et al. 2012b Pomacentrus amboiensis 700, 850 μatm 440 μatm 850 μatm 4d: settlement Visual social Learning Forsgren et al. 2013 Gobiusculus flavescens 1400 μatm 370 μatm 1400 μatm 9-12d; NH* Phototaxis

Lönnstedt et al. 2013 Pomacentrus amboiensis 880 μatm 440 μatm 880 μatm 4 d; settlement Vision/Chemosensory

Chung et al. 2014 Acanthochromis polyacanthus 944 μatm 466 μatm 944 μatm 6-7d; small Retinal function 1 33 Vision/Predation Spady et al. 2014 Idiosepius pygmaeus 626, 956 μatm 447 μatm 626 μatm 5d:adult threat

The preference of larval fish to swim towards a light source (phototaxis) is essential for feeding and survival (Karlsen and Mangor-Jensen 2001). Under elevated

CO2 conditions, newly-hatched larvae of a temperate goby (Gobiusculus flavescens) experienced a 65-70% increase in positive phototaxis, both in terms of proportion of larvae to reach the light source in 2 minutes and the speed of the fastest larvae per trial (Forsgren et al. 2013). Although positive phototaxis is beneficial to larvae, an increase this drastic suggests visual hypersensitivity due to an overload of brain activity, which could be maladaptive (Forsgren et al. 2013). Anti-predator behavior is crucial to the survival of marine organisms and can include reduced foraging, increased shelter use and “bobbing” signaling behavior when threatened. Appropriate anti-predator behavior can be innate or learned through visual social cues (Kelley and Magurran 2003). When exposed to visual cues from a predator, CO2-treated juvenile damselfish, Pomacentrus amboinensis, displayed weaken anti-predator behavior by continued foraging, lack of bobbing behavior, and normal activity levels (Ferrari et al. 2012b). Moreover, P. amboinensis exposed to ocean acidification conditions were unable to socially learn appropriate anti-predator behavior from visual cues from conspecifics (Ferrari et al. 2012a). Anti-predatory behavior has also been tested in the invertebrate tropical squid,

Idiosepius pygmaeus. When presented with a visual threat stimulus, squid exposed to elevated CO2 retained their predator escape ability (no difference in reaction time or distance moved), but the choice of escape behavior shifted to jet propulsion over defensive arm positioning (Spady et al. 2014). While this study shows critical behavioral changes to a visual threat, it suggests a neural alteration rather than sensory impairment.

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From the few studies so far conducted, it appears that anti-predatory responses to visual cues are affected but not entirely lost (Ferrari et al. 2012a,b; Spady et al. 2014). In contrast, the response to olfactory cues can be completely reversed (Dixson et al. 2010). Visual perception may provide sensory compensation for the loss or alteration of chemosensory perception (Lonnstedt et al. 2013; Cripps et al. 2011; Devine et al. 2012a). A possible mechanism for altered behaviors in response to visual cues may be slowed retinal function, characterized by critical clicker fusion (CFF) (Chung et al. 2014). CFF is the threshold at which an animal perceives a flickering light as continuous and is variable depending on lifestyle and environmental illumination (Horodysky et al. 2008; Smolka et al. 2013). The CFF of damselfish, Acanthochromis polyacanthus, after exposure to ocean acidification conditions was significantly reduced compared to controls. A decrease in CFF could weaken the ability to react to fast events during foraging and predator avoidance. This decrease in CFF was negated after exposure to the GABAA antagonist, gabazine, suggesting an underlying influence of neurotransmitter overactivation (Nilsson et al. 2012; see mechanisms below).

B.5 Other senses

Migrating marine animals are thought to rely on a magnetic sense for navigation using the Earth’s magnetic field or anomalies within the Earth’s crust at mid-ocean ridges (Walker et al. 2002). The ability to discriminate between magnetic fields has been shown in teleosts (Quinn 1980), elasmobranchs (Kalmijn 1978), spiny lobsters (Lohmann et al. 1995) and sea turtles (Lohmann and Lohmann 1996). The precise mechanism by which marine organisms detect magnetic fields is unknown,

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although multiple hypotheses have been proposed (Lohman and Johnsen 2000). While no definitive studies have been conducted, it is plausible to hypothesize that the magnetic sense in marine organisms would be influenced by ocean acidification due to neural impairment. The reception of the magnetic field cues may not be affected, but the interpretation of these cues may be influenced, leading to complications with navigation. The lateral line sense is highly conserved among aquatic vertebrates including fishes and amphibians and allows for the detection of water currents and pressure gradients (Bleckmann and Zelick 2009). The lateral line system is composed of a network of mechanoreceptors, or neuromasts, that reside in canals over the head and along the side of the organisms (Flock and Wersall 1962) and plays a critical role in rheotaxis, or the orientation into an oncoming current (Montgomery et al. 1997).

Currently, no studies have been conducted on the effects of elevated CO2 on the effectiveness of the lateral line sensory system. We hypothesize that the perception of external cues through the lateral line system (pressure, current direction) will be influenced due to the neural component of sensory perception and advocate for research to test this hypothesis. The lateral line system has evolved specialized functions in sharks, skates and rays, which have the unique ability to detect bioelectric stimuli using ampullary electrosense (Kalmijn 1972). Elasmobranchs use this sense during foraging, social interactions, reproduction and anti-predatory behaviors (Wilkens and Hofmann 2005). Structurally, elasmobranchs are equipped with electroreception organs called ampullae of Lorenzini, clusters of which can detect not only small local electric fields emanating from prey, but also uniform electric fields from background structure

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(Sisneros and Tricas 2002). Similar to chemoreceptors, electroreceptors are located on or directly connected to the surface of the organism, possibly making them more vulnerable to changing ocean chemistry. No studies to date have assessed the effect of ocean acidification on electrosense. However it is reasonable to assume changes will occur for two reasons: (1) changes in salt concentration, temperature and pH are known to influence the conductivity of bioelectric fields and (2) similar to other senses, the endpoint of electroreception is in neural processing and therefore elasmobranchs may experience alterations in how they perceive electrical pulses.

B.6 Mechanisms

Several physiologic mechanisms have been proposed to explain the disruption of sensory perception due to elevated CO2. Briffa et al. (2012) highlight possible mechanisms for info-disruption in aquatic organisms. Chemoreception relies on the connection between sensory receptor molecules and odor molecules. A change in seawater chemistry may shift the ionic charge of the receptor sites (Tierney and Atema 1987) or the chemical molecules (Hara 1976; Brown et al. 2002; Leduc et al. 2004). While this has mainly been tested in freshwater systems, which experience different chemical environments, it may be a potential mechanism of sensory impairment in the marine system. In fact, Roggatz et al. (2016) discovered that an acidified ocean could modify the charge, shape and hydrophobicity on peptide signaling cues, thereby affecting receptor-cue interactions and requiring a higher cue concentration to induce a response. On the other hand, sensory organs as a whole may be affected by elevated

CO2, however a connection between structural alterations and sensory impairment has

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not been observed thus far for chemoreception (Munday et al. 2009; de la Haye et al. 2012) or sound (Simpson et al. 2011). Finally, evidence including multiple sensory system impairment, cognitive alterations involving lateralization (Domenici et al. 2011) and learning (Ferrari et al.

2012b), and the observation that high CO2 treated fish received chemical cues but responded inappropriately, suggests a disruption in processing at the brain or neural level.

The GABAA receptor is the primary neurotransmitter receptor in the vertebrate brain (Bormann et al 1987), and many invertebrates have GABAA-like receptors (Hutton et al. 1993; Lunt 1991). These receptors are vital to multiple sensory systems across diverse taxa (ex: Pierobon et al. 2004; Igarashi et al. 2009; Yang et al. 2004).

Nilsson et al. (2012) revealed that the inhibitory neurotransmitter receptor, GABAA, plays a role in the chemosensory and lateralization changes observed in reef fish. Acid-base balance allows fish to maintain functional extracellular pH through the

- - adjustment of blood plasma HCO3 and Cl . The ability to maintain appropriate intercellular pH is critical to cell function (Putnam and Roos 1997) and can be linked to metabolism and iono-regulation (Widdicombe and Spicer 2008). In ambient

- - seawater conditions, the ion gradients allow an influx of Cl and HCO3 through the neuronal membrane, which leads to hyperpolarization and inhibition of the neuron

(Heuer and Grosell 2014). When held in elevated pCO2, fish control their acid-base

- - balance by accumulating HCO3 and ejecting Cl , which stabilizes the intracellular pH

- but increases the concentration of HCO3 and pCO2 in the extracellular fluid (Nilsson et al. 2012). The expulsion of Cl- changes the ion gradients, resulting in the extrusion rather than the incursion of these ions when GABA binds to GABAA receptors.

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Therefore, the membrane switches from a hyperpolarizing, inhibitory current to a depolarizing, excitatory current. Fish treated with gabazine, a GABAA antagonist, returned to normal olfactory and lateralization behaviors (Nilsson et al. 2012). This mechanism has been confirmed in retinal function and learning in damselfish (Chung et al. 2014; Chivers et al. 2014), predator evasion in a gastropod (Watson et al. 2014), lateralization in three-spined stickleback (Lai et al. 2015) and anxiety in juvenile salmon (Ou et al. 2015) and rockfish (Hamilton et al. 2014). In a novel approach,

Regan et al. (2016) observed the effect of low CO2 exposure to a fish native to hypercapnic waters (Pangasianodon hypophthalmus) and found behavioral alterations that were repaired by the addition of gabazine.

B.7 Comparison between taxa

An understanding of the sensory systems affected by ocean acidification and the mechanisms behind this impairment can guide our predictions of which marine taxa are at greater risk of sensory impairment. As mentioned above, the excitation of

GABAA receptors as a side effect of acid-base regulation is thought to be a primary mechanism for sensory impairment. Therefore we might expect organisms with partial or complete acid-base regulation to be more susceptible to altered sensory processing (Nilsson et al. 2012). The ability to regulate acid-base compensation can be a function of metabolic rate, metabolic rate fluctuations, current environmental hypercapnia, and ontogenetic hypercapnia (Melzner et al. 2009). High-power organisms or those that experience long developmental periods within their egg, such as fish, cephalopods and some crustaceans, as well as organisms that experience elevated pCO2 levels (intertidal zones, estuaries, upwelling areas, etc) typically exhibit complete or near

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complete extracellular compensation. Conversely, organisms characterized by hypometabolism and primarily pelagic development such as bivalves, echinoderms and sipunculid worms, as well as those located in stable environmental conditions such as the deep sea typically show incomplete or inferior compensation (Meltzner et al. 2009; Widdicombe and Spicer 2008). However, the current review shows modified sensory perception in fish (elasmobranchs and teleosts), gastropods, crustaceans, echinoderms and cnidarians, a pattern that does not match the extent of acid-base regulation. Beyond relative completion of acid-base regulation, extent of sensory impairment may reflect adaptation or acclimation potential. Numerous studies have shown transgenerational acclimation of physiological or metabolic alterations under high CO2 conditions (Miller et al. 2012; Murray et al. 2014; Ho and Berggren 2012). However, transgenerational plasticity is absent or partial in studies assessing behavior or sensory function (Allan et al. 2014; Welch et al. 2014). In general, adaptational differences may be evident when life span and reproductive strategies are compared, with shorter-lived r-selected species able to more readily pass on epigenetic plasticities than longer-lived k-selected organisms.

Finally, taxa differences in sensory impairment in a high CO2 ocean may reflect the ecology of the organism in regards to relative use of sensory systems. While antipredatory responses to visual cues appear to be weakened by ocean acidification (Ferrari et al. 2012b; Lonnstedt et al. 2013), they are not completely reversed or lost as in chemosensory cues (Dixson et al. 2010; Lonnstedt et al. 2013). This suggests that more visually-dependent organisms may be better equipped to handle ocean acidification conditions. Further CO2-exposed damselfish have altered responses to

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predatory (Lonnstedt et al. 2013) and habitat (Devine et al. 2012a) chemosensory cues, however, these effects are nonexistent when multiple cues were presented, suggesting sensory compensation. Therefore, organisms with multiple different sensory systems may be more equipped to perceive their environment with the loss or impairment of one, compared with simpler organisms that lack multiple senses, therefore the loss of critical senses are felt more soundly.

B.8 Future directions

This review highlights the current knowledge on the effects of ocean acidification (Fig.1). Synthesizing this field has revealed trends, biases, and a lack of standardized methodology that, if rectified, would significantly improve our understanding of the full extent of elevated CO2 on sensory systems.

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Neural functioning - Chemosensory (Table 1)

Sensory impairment at the central processing level (GABAA receptors) Known effects: altered perception of Evidence: Multiple senses affected, Incorrect homing, settlement, predator/prey cues responses rather than lack of response, additional Tested organisms: Reef and temperate Lish, cognitive alterations including lateralization and learning coral larvae, gastropods, sea urchins, and crusteaceans

Vision (Table 3)

Known effects: Altered phototaxis sensitivity, visual predator detection and response, visual learning, retinal function Audition (Table 2) Tested organisms: Reef Lish, temperate goby, squid

Known effects: Altered perception of auditory homing cues, increased otolith size Other senses: Tested organisms: Temperate and tropical Lateral Line system, Lish Electrosense, Magnetic sense Evidence for impairment: No current studies, but alterations expected due to processing at neural level Figure B.1: Infogram depicting the summarized impacts of ocean acidification on the sensory systems of a model vertebrates and invertebrate organisms.

Notable variations in the CO2 level tested, length of exposure and habituation period has made species or systems comparisons difficult if not impossible (Table A1-

A3). Variations in CO2 levels tested are expected and acceptable as long as these levels reflect the projected levels expected in that particular region. However, using global averages for ocean acidification may not be the most accurate prediction for specific regions, thus local and regional future pH estimates should be determined before experimentation. Habituation period is critical at the start of experiments as acid-base regulation can begin within 15 minutes, with full compensation within a few hours in at least one species (Espaugh et al. 2012). There is no established

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standardized methodology for habituation length or exposure time, both of which can have substantial influence on results and should be normalized. Just as the synthesis of current studies revealed high variability, it also reveals the biases in organisms and species studied. The majority of studies focus on fishes, larvae or juveniles, reef ecosystems, and chemosensory perception. Expansion of this field involves (1) researching the diversity of responses exhibited by marine invertebrates and non-fish vertebrates, (2) tracking individual responses through ontogeny and into adulthood to understand the influence of life stage, (3) focusing on other ecosystems that may be even more threatened by ocean acidification than reefs, including estuaries (Meltzner et al. 2013) and upwelling zones (Steckbauer et al. 2015) because of the additive effect of hypoxia on hypercapnia response (Melzner et al. 2013), and (4) designing behavioral or physiological assays to test neglected sensory systems such as electrosense, magnetic perception and the lateral line system. As the field advances, focus should shift towards creating more realistic conditions while testing sensory perception. Realistically, marine ecosystems will face multiple stressors in concordance with ocean acidification, including temperature, pollution, hypoxia and habitat degradation. A plethora of studies have investigated the combined effects of temperature and lowered pH on calcification, survival and growth

(Harvey et al. 2013; Lefevre 2016), but few have addressed behavior or sensory impairment. Similarly, physiological studies often address both acidification and hypoxia (Pörtner 2008; Meltzner et al. 2013), but this has yet to be assessed for behavior. Additionally, an effort to produce more realistic studies will incorporate natural pH variation into laboratory design. Often, the nightly decrease in pH from respiration is lower than the oceanic pH expected in the next century (Hofmann et al.

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2011), yet few researchers have included these fluctuations in their experimental design. Natural variations in environmental pH are predicted to expedite the onset of elevated CO2 thresholds (Shaw et al. 2013a) and the magnitude of variation may increase non-linearly due to a weakening of the environmental buffering system (Shaw et al. 2013b). However, exposure to variable environmental pH conditions can reduce sensitivity to manipulated pH conditions at least in terms of survival (Lewis et al. 2013), but this has yet to be tested in terms of sensory impairment. Further research focusing on acclimation potential would also be critical since ocean acidification is expected to occur gradually throughout the next century (see taxa comparison section). Finally, more realistic studies must scale-up to determine community-level effects. This can begin by testing multiple species in an interaction; since both partner in predator-prey interactions have the potential to experience sensory impairment in future conditions, studies investigating both sides simultaneously could provide valuable information beyond individual species. Mesocosm experiments are an obvious next step and have already been conducted when testing for survival, bleaching and community structure changes (ref). Some work has shown increase mortality due to high CO2 behavioral or sensory alterations after transporting treated fish back to the natural environment (Munday et al. 2010). CO2 seeps in Papua New Guinea can act as an ecologically relevant natural laboratory, where community-scale effects of sensory impairment can be observed without typical experimental caveats including habituation, shock potential and handling stress. Previous work has shown that fish from the natural seeps exhibited the same altered behaviors as seen in the laboratory, although there was no difference in community structure (Munday et al.

2014). Natural CO2 seeps are close predictors of future conditions, yet still retain the

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ability to act as sinks to allochthonous larvae, therefore results should be considered with caution. Ultimately as the field advances, effective and meaningful studies must strive for realistic studies incorporating whole-community dynamics, whether through multiple species interactions in the laboratory setting, mesocosm experiments, or the use of natural CO2 environments.

B.9 Conclusions

Ocean acidification will have a profound effect on the sensory perception of marine organisms, with highly variable responses based on species, life-stage, ecosystem, and parental influence. The field has overwhelmingly been dominated by studies of coral reef vertebrate larvae, and this significant bias needs to be addressed and corrected if we want to develop a more accurate picture of future ocean ecosystems. Further studies should focus on (1) expanding the range of species studied, (2) including marine ecosystems outside of coral reefs, especially estuaries and mangroves, (3) assessing the potential of transgenerational acclimation or genetic adaptation to elevated CO2 levels, (4) emphasizing more realistic laboratory experiments to include multiple stressors and environmental variability, and (5) scaling up to reveal community-wide effects of sensory and behavioral adjustments to increased CO2. This diversification of ecosystem perspectives will expand our understanding of sensory impairment under ocean acidification conditions.

B.10 References

Ache BW, Young JM. 2005. Olfaction: diverse species, conserved principles. Neuron. 48:417-430.

145

Albright R, Mason B, Miller M, Langdon C. 2010. Ocean acidification compromises recruitment success of the threatened Caribbean coral Acropora palmata. Proc Natl Acad Sci USA. 107:20400-20404.

Albright R, Langdon C. 2011. Ocean acidification impacts multiple early life history processes of the Caribbean coral Porites astreoides. Global Change Biol. 17:2478-2487.

Allan BJM, Miller GM, McCormick MI, Domenici P, Munday PL. 2014. Parental effects improve escape performance of juvenile reef fish in a high-CO2 world. Proc Biol Sci. 281:20132179.

American Heritage Medical Dictionary. 2007. Sensory input (n.d.). Retrieved March 5 2017 from http://medical-dictionary.thefreedictionary.com/sensory+input

Arendt D. 2003. Evolution of eyes and photoreceptor cell types. Int J Dev Biol. 47:563-571.

Arshavsky VY, Lamb TD, Pugh Jr. EN. 2002. G proteins and phototransduction. Annu Rev Physiol. 64:153-187.

Atema J, Kingsford MJ, Gerlach G. 2002. Larval reef fish could use odor for detection, retention and orientation to reefs. Mar Ecol Prog Ser. 242:151-160.

Barry JP, Lovera C, Buck KR, Peltzer ET, Taylor JR, Walz, P, Whaling PJ, Brewer PG. 2014. Use of a Free Ocean CO2 Enrichment (FOCE) system to evaluate the effects of ocean acidification on the foraging behavior of a deep-sea urchin. Environ Sci Technol. 48:9890-9897.

Bignami S, Enochs IC, Manzello DP, Sponaugle S, Cowen RK. 2013a. Ocean acidification alters the otoliths of a pantropical fish species with implications for sensory function. Proc Natl Acad Sci USA. 110:7366-7370.

Bignami S, Sponaugle S, Cowen RK. 2013b. Response to ocean acidification in larvae of a large tropical marine fish, Rachycentron canadum. Global Change Biol. 19:996-1006.

Bleckmann, H, Zelick R. 2009. Lateral line system of fish. Integr Zool. 4:13-25.

Bormann J, Hamill OP, Sakmann B. 1987. Mechanism of anion permeation through channels gated by glycine and γ-aminobutyric acid in mouse cultured spinal neurones. J Physiol. 385:243-286.

Briffa M, de la Haye K, Munday PL. 2012. High CO2 and marine animal behaviour: Potential mechanisms and ecological consequences. Mar Pullut Bull. 65:1519- 1528.

146

Brown GE, Adrian JC, Lewis MG, Tower JM. 2002. The effects of reduced pH on chemical alarm signaling in ostariophysan fishes. Can J Fish Aquat Sci. 59:1331–1338.

Buck L, Axel R. 1991. A novel multigene family may encode odorant receptors: a molecular basis for odor recognition. Cell. 65:175-187.

Budelmann BU. 1992. Hearing in nonarthropod invertebrates. In: The evolutionary biology of hearing (pp. 141-155). Springer New York.

Byrne M. 2011. Impact of ocean warming and ocean acidification on marine invertebrate life history stages: vulnerabilities and potential for persistence in a changing ocean. Oceanogr Mar Biol Annu Rev 49:1-42.

Caley MJ, Carr MH, Hixon MA, Hughes TP, Jones GP, Menge BA. 1996. Recruitment and the local dynamics of open marine populations. Annu Rev Ecol Evol Syst. 27:477-500.

Checkley DM, Dickson AG, Takahashi M, Radich JA, Eisenkolb N, Asch R. 2009. Elevated CO2 enhances otolith growth in young fish. Science. 324:1683.

Chivers DP, McCormick MI, Nilsson GE, Munday PL, Watson S-A, Meekan MG, Mitchell MD, Corkill KC, Ferrari MCO. 2014. Impaired learning of predators and lower prey survival under elevated CO2: a consequence of neurotransmitter interference. Global Change Biol. 20:515-522.

Chung W, Marshall NJ, Watson S, Munday PL, Nilsson GE (2014) Ocean acidification slows retinal function in a damselfish through interference with GABAA receptors. J Exp Biol. 217:323-326.

Clements JC, Hunt HL. 2015. Marine animal behaviour in a high CO2 ocean. Mar Ecol Prog Ser. 536:259-279.

Cole KS, Shapiro DY. 1995. Social facilitation and sensory mediation of adult sex change in a cryptic, benthic marine goby. J Exp Mar Biol Ecol. 186:65-75.

Cripps IL, Munday PL, McCormick MI. 2011. Ocean acidification affects prey detection by a predatory reef fish. Plos One. 6:e22736. de la Haye KL, Spicer JI, Widdicombe S, Briffa M. 2012. Reduced pH sea water disrupts chemo-responsive behaviour in an intertidal crustacean. J Exp Mar Biol Ecol. 412:134-140.

Derby CD. 1982. Structure and function of cuticular sensilla of the lobster, Homarus americanus. J Crust Biol. 2:1-21.

147

Devine BM, Munday PL. 2013. Habitat preferences of coral-associated fishes are altered by short-term exposure to elevated CO2. Mar Biol. 160:1955-1962.

Devine BM, Munday PL, Jones GP. 2012a. Rising CO2 concentrations affect settlement behavior of larval damselfishes. Coral Reefs. 31:229-238.

Devine BM, Munday PL, Jones GP. 2012b. Homing ability of adult cardinalfish is affected by elevated carbon dioxide. Oecologia. 168:269-276.

Dixson DL, Munday PL, Jones GP. 2010. Ocean acidification disrupts the innate ability of fish to detect predator olfactory cues. Ecol Lett. 13: 68-75.

Dixson DL, Jones GP, Munday PL, Planes S, Pratchett MS, Srinivasan M, Syms C, Thorrold SR. 2008. Coral reef fish smell leaves to find island homes. P Roy Soc Lond B Bio. 275:2831-2839.

Dixson DL, Abrego D, Hay ME. 2014. Chemically mediated behavior of recruiting corals and fishes: A tipping point that may limit reef recovery. Science. 245:892-897.

Dixson DL, Jennings AR, Atema J, Munday PL. 2015. Odor tracking in sharks is reduced under future ocean acidification conditions. Global Change Biol. 21:1454-1462.

Domenici P, Allan B, McCormick MI, Munday PL. 2011. Elevated carbon dioxide affects behavioural lateralization in a coral reef fish. Biol Lett. 8:78-81.

Doney SC, Fabry VJ, Feely RA, Kleypas JA. 2009. Ocean acidification: the other CO2 problem. Ann Rev Mar Sci. 1:169-192.

Doropoulos C, Ward S, Diaz‐Pulido G, Hoegh‐Guldberg O, Mumby PJ. 2012. Ocean acidification reduces coral recruitment by disrupting intimate larval‐algal settlement interactions. Ecol Lett. 15:338-346.

Esbaugh AJ, Hauer R, Grosell M. 2012. Impacts of ocean acidification on respiratory gas exchange and acid0base balance in a marine telost, Opsanus beta. J Comp Physiol B. 184:921-934.

Fabry VJ, Seibel BA, Feely RA, Orr JC. 2008. Impacts of ocean acidification on marine fauna and ecosystem processes. ICES J Mar Sci: Journal du Conseil: 414-432.

Ferrari MC, McCormick MI, Munday PL, Meekan MG, Dixson DL, Lönnstedt O, Chivers DP. 2012a. Effects of ocean acidification on visual risk assessment in coral reef fishes. Funct Ecol. 26:553-558.

148

Ferrari MCO, Manassa RP, Dixson DL, Munday PL, McCormick MI, Meekan MG, Sih A, Chivers DP. 2012b. Effects of ocean acidification on learning in coral reef fishes. PloS One. 7:e31478.

Ferrari MCO, Dixson DL, Munday PL, McCormick MI, Meekan MG, Sih A, Chivers DP. 2011a. Intrageneric variation in antipredator responses of coral reef fishes affected by ocean acidification: implications for climate change projections on marine communities. Global Change Biol. 17:2980-2986.

Ferrari MCO, Wisenden BD, Chivers DP. 2010. Chemical ecology of predator-prey interactions in aquatic ecosystems: a review and prospectus. Can J Zool. 88:698–724.

Flock Å, Wersäll J. 1962. A study of the orientation of the sensory hairs of the receptor cells in the lateral line organ of fish, with special reference to the function of the receptors. J Cell Biol. 15:19-27.

Forsgren E, Dupont S, Jutfelt F, Amundsen T. 2013. Elevated CO2 affects embryonic development and larval phototaxis in a temperate marine fish. Ecol Evol. 3:3637-3646.

Franke A, Clemmesen C. 2011. Effect of ocean acidification on early life stages of Atlantic herring (Clupea harengus L.). Biogeosciences. 8:3697-3707.

Frommel AY, Schubert A, Piatkowski U, Clemmesen C. 2013. Egg and early larval stages of Baltic cod, Gadus morhua, are robust to high levels of ocean acidification. Mar Biol. 160:1825-1834.

Gagliano M, Depczynski M, Simpson SD, Moore JA. 2008. Dispersal without errors: symmetrical ears tune into the right frequency for survival. P Roy Soc Lond B Bio. 275:527-534.

Hamilton TJ, Holcombe A, Tresguerres M. 2014. CO2-induced ocean acidification increases anxiety in Rockfish via alteration of GABAA receptor functioning. Proc R Soc B. 281:20132509.

Hara TJ. 1994. Olfaction and Gustation in fish: an overview. Acta Physiol Scand. 152:207-217

Hara T. 1976. Effects of pH on the olfactory responses to amino acids in rainbow trout Salmo gairdneri. Comp Biochem Physiol. A 54:37–39.

Hay ME. 2009. Marine chemical ecology: chemical signals and cues structure marine populations, communities, and ecosystems. Ann Rev Mar Sci. 1:193-212

149

Heenan A, Simpson SD, Braithwaite VA. 2009. Testing the generality of acoustic cue use at settlement in larval coral reef fish. In Proc. of the 11th Int. Coral Reef Symp., Fort Lauderdale, Florida, 7–11 July 2008, Session 16, pp. 554–558.

Heuer RM, Grosell M. 2014. Physiological impacts of elevated carbon dioxide and ocean acidification on fish. Am J Physiol Regul Integr Comp Physiol. 307:R1061-R1084.

Ho DH, Burggren WW. 2012. Parental hypoxic exposure confers offspring hypoxia resistance in zebrafish (Danio rerio). J Exp Biol. 215:4208-16

Hofmann GE, Smith JE, Johnson KS, Send U, Levin LA, Micheli F, Paytan A, Price NN, Peterson B, Takeshita Y, et al. 2011. High-frequency dynamics of ocean pH: a multi-ecosystem comparison. PloS one. 6:e28983.

Horodysky AZ, Brill RW, Warrant EJ, Musick JA, Latour RJ. 2008. Comparative visual function in five sciaenid fishes inhabiting Chesapeake Bay. J Exp Biol. 211:3601-3612.

Hurst TP, Fernandez ER, Mathis JT, Miller JA, Stinson CM, Ahgeak EF. 2012. Resiliency of juvenile walleye Pollock to projected levels of ocean acidification. Aquat Biol. 17:247-259.

Hutton ML, Harvey RJ, Earley FG, Barnard EA, Darlison MG. 1993. A novel invertebrate GABAA receptor-like polypeptide sequence and pattern of gene expression. FEBS Lett. 326:112-116.

Igarashi A, Zadzilka N, Shirahata M. 2009. Benzodiazepines and GABA-GABAA receptor system in the cat carotid body. In Arterial Chemoreceptors. pp. 169- 175. Springer Netherlands.

Jeffs A, Tolimieri N, Montgomery JC. 2003. Crabs on cue for the coast: the use of underwater sound for orientation by pelagic crab stages. Mar Freshwater Res. 54:841-845.

Jutfelt F, Hedgärde M. 2013. Atlantic cod actively avoid CO2 and predator odor, even after long-term CO2 exposure. Front Zool. 10:1-8.

Kaifu K, Akamatsu T, Segawa S. 2008. Underwater sound detection by cephalopod statocyst. Fisheries Sci. 74:781-786.

Kalmijn AJ. 1978. Experimental evidence of geomagnetic orientation in elasmobranch fishes. In: Animal migration, navigation, and homing (pp. 347-353). Springer Berlin Heidelberg.

150

Kalmijn AJ. 1972. Bioelectric fields in sea water and the function of the ampullae of Lorenzini in elasmobranch fishes. Scripps Institution of Oceanography.

Karlsen Ø, Mangor-Jensen A. 2001. A correlation between photoactic response and first-feeding of Atlantic halibut (Hippoglossus hippoglossus L.) larvae. Aquac Res. 32:907-912.

Kasumyan AO. 2004. The olfactory system in fish: structure, function, and role in behavior. J Ichthyol. 44:S180-S223.

Kelley JL, Magurran AE. 2003. Learned predator recognition and antipredator responses in fishes. Fish Fish. 4:216-226.

Kroeker KJ, Kordas RL, Crim RN, Singh GG. 2010. Meta‐analysis reveals negative yet variable effects of ocean acidification on marine organisms. Ecol Lett. 13:1419-1434.

Kroeker KJ, Sanford E, Jellison BM, Gaylord B. 2014. Predicting the effects of ocean acidification on predator-prey interactions: a conceptual framework based on coastal molluscs. Biol Bull. 226:211-222.

Lai F, Jutfelt F, Nilsson GE. 2015. Altered neurotransmitter function in CO2-exposed stickleback (Gasterosteus aculeatus): a temperate model species for ocean acidification research. Conserv Physiol. 3:cov018.

Land MF, Fernald RD. 1992. The evolution of eyes. Annu Rev Neurosci. 15:1-29.

Land MF, Nilsson D-E. 2002. Animal eyes. Oxford: Oxford University Press.

Lecchini D, Shima J, Banaigs B, Galzin R. 2005. Larval sensory abilities and mechanisms of habitat selection of a coral reef fish during settlement. Oecologia. 143:326-334.

Leduc AOHC, Kelly JM, Brown GE. 2004. Detection of conspecific alarm cues by juvenile salmonids under neutral and weakly acidic conditions: laboratory and field tests. Oecologia. 139:318–324.

Lefevre S. 2016. Are global warming and ocean acidification conspiring against marine ectotherms? A meta-analysis of the respiratory effects of elevated temperature, high CO2 and their interaction. Conserv Physiol. 4: cow009.

Leis JM, Carson-Ewart BM, Cato DH. 2002. Sound detection in situ by the larvae of a coral-reef damselfish (Pomacentridae). Mar Ecol Prog Ser. 232:259-268.

151

Leis JM, Sweatman HPA, Reader SE. 1996. What the pelagic stages of coral reef fishes are doing out in blue water: daytime field observations of larval behavioural capabilities. Mar Freshwater Res. 47:401–411.

Lewis CN, Brown KA, Edwards LA, Cooper G, Findlay HS. 2013. Sensitivity to ocean acidification parallels natural pCO2 gradients experienced by Arctic copepods under winter sea ice. Proc Nat Acad Sci USA. 110:E4960-E4967.

Lima SL, Dill LM. 1990. Behavioral decisions made under the risk of predation: a review and prospectus. Can J Zool. 68:619-640.

Lohmann K, Pentcheff N, Nevitt G, Stetten G, Zimmer-Faust R, Jarrard H, Boles LC. 1995. Magnetic orientation of spiny lobsters in the ocean: experiments with undersea coil systems. J Exp Biol. 198:2041-2048.

Lohmann K, Lohmann C. 1996. Orientation and open-sea navigation in sea turtles. J Exp Biol. 199:73-81.

Lohmann KJ, Johnsen S. 2000. The neurobiology of magnetoreception in vertebrate animals. Trends Neurosci. 23:153-159.

Lönnstedt OM, Munday PL, McCormick MI, Ferrari MCO, Chivers DP. 2013. Ocean acidification and responses to predators: can sensory redundancy reduce the apparent impacts of elevated CO2 on fish? Ecol Evol. 3:3565-3575.

Lovell JM, Findlay MM, Moate RM, Yan HY. 2005. The hearing abilities of the prawn, Palaemon serratus. Comp Biochem Phys A. 140:89-100.

Lunt GG. 1991. GABA and GABA receptors in invertebrates. Sem Neurosci. 3:251- 258.

Lychakov DV, Rebane YT. 2000. Otolith regularities. Hear Res. 143:83-102.

Maneja RH, Frommel AY, Geffen AJ, Folkvord A, Piatkowski U, Chang MY, Clemmesen C. 2013. Effects of ocean acidification on the calcification of otoliths of larval Atlantic cod Gadus morhua. Mar Ecol Prog Ser. 477: 251- 258.

Manríquez PH, Jara ME, Mardones ML, Torres R, Navarro JM, Lardies MA, Vargas CA, Duarte C, Lagos NA. 2014. Ocean acidification affects predator avoidance behavior but not prey detection in the early ontogeny of a keystone species. Mar Ecol Prog Ser. 502:157-167.

152

Melzner F, Gutowska MA, Langenbuch M, Dupont S, Lucassen M, Thorndyke MC, Bleich M, Pörtner HO. 2009. Physiological basis for high CO2 tolerance in marine ectothermic animals: pre-adaptation through lifestyle and ontogeny?. Biogeosciences. 6:2313-2331.

Melzner F, Thomsen J, Koeve W, Oschlies A, Gutowska MA, Bange HW, Hansen HP, Körtzinger A. 2013. Future ocean acidification will be amplified by hypoxia in coastal habitats. Mar Biol. 160:1875-1888.

Miller GM, Watson SA, Donelson JM, McCormick MI, Munday PL. 2012. Parental environment mediates impacts of increased carbon dioxide on a coral reef fish. Nat Clim Change. 2:858-861.

Montgomery JC, Baker CF, Carton AG. 1997. The lateral line can mediate rheotaxis in fish. Nature. 389:960-963.

Munday PL, Dixson DL, Donelson JM, Jones GP, Pratchett MS, Devitsina, GV, Døving KB. 2009. Ocean acidification impairs olfactory discrimination and homing ability of a marine fish. P Natl Acad Sci USA. 106:1848-1852.

Munday PL, Dixson DL, McCormick MI, Meekan M, Ferrari MC, Chivers DP. 2010. Replenishment of fish populations is threatened by ocean acidification. P Natl Acad Sci USA. 107:12930-12934.

Munday PL, Pratchett MS, Dixson DL, Donelson JM, Endo GGK, Reynolds AD, Knuckey R. 2013. Elevated CO2 affects the behavior of an ecologically and economically important coral reef fish. Mar Biol. 160:2137-2144.

Munday PL, Cheal AJ, Dixson DL, Rummer JL, Fabricius KE. 2014. Behavioural impairment in reef fishes caused by ocean acidification at CO2 seeps. Nat Clim Change. 4:487-492.

Munday PL, Hernaman V, Dixson DL, Thorrold SR. 2011a. Effect of ocean acidification on otolith development in larvae of a tropical marine fish. Biogeosciences. 8:1631-1641.

Munday PL, Gagliano M, Donelson JM, Dixson DL, Thorrold SR. 2011b. Ocean acidification does not affect the early life history development of a tropical marine fish. Mar Ecol Prog Ser. 423:211-221.

Montgomery JC, Baker CF, Carton AG. 1997. The lateral line can mediate rheotaxis in fish. Nature. 389:960-963.

Murray CS, Malvezzi A, Gobler CJ, Baumann H. 2014. Offspring sensitivity to ocean acidification changes seasonally in a coastal marine fish. Mar Ecol Prog Ser. 504:1-11.

153

Myrberg Jr AA. 1997. Underwater sound: its relevance to behavioral functions among fishes and marine mammals. Mar Freshw Behav Phy. 29:3-21.

Nagelkerken I, Munday PL. 2016. Animal begabiour shapes the ecological effects of ocean acidification and warming: moving from individual to community-level responses. Global Change Biol. 22:974-989.

Nilsson GE, Dixson DL, Domenici P, McCormick MI, Sørensen C, Watson SA, Munday PL. 2012. Near-future carbon dioxide levels alter fish behaviour by interfering with neurotransmitter function. Nat Clim Change. 2:201-204.

Ou M, Hamilton TJ, Eom J, Lyall EM, Gallup J, Jiang A, Lee J, Close DA, Yun SS, Brauner CJ. 2015. Responses of pink salmon to CO2-induced aquatic acidification. Nat Clim Change. 5:950-955.

Pawlik JR. 1992. Chemical ecology of the settlement of benthic marine invertebrates. Oceanogr Mar Biol Annu Rev. 30:273-335.

Perry DM, Redman DH, Widman JC, Meseck S, King A, Pereira JJ. 2015. Effect of ocean acidification on growth and otolith condition of juvenile scup, Stenotomus chrysops. Ecol Evol. 5:4187-4196.

Pierobon P, Tino A, Minei R, Marino G. 2004. Different roles of GABA and glycine in the modulation of chemosensory responses in Hydra vulgaris (Cnidaria, Hydrozoa). In Coelenterate Biology. pp. 59-66. Springer Netherlands.

Polese G, Bertapelle C, Di Cosmo A. 2016. Olfactory organ of Octopus vulgaris: morphology, plasticity, turnover and sensory characterization. Biol Open. 017764.

Popper AN, Fay RR. 1973. Sound detection and processing by teleost fishes: a critical review. J Acoust Soc Am. 53:1515-1529.

Pörtner H-O. 2008. Ecosystem effects of ocean acidification in times of ocean warming: a physiologist’s view. Mar Ecol Prog Ser. 373:203-217.

Pumphrey RJ. 1950. Hearing. In: Symposia of the Society for Experimental Biology (Vol. 4, pp. 3-18). Univ Cambridge Dept Zoology, Downing St, Cambridge CB2 3EJ, Cambs, England: Company Biologists LTD.

Putnam RW, Roos A. 1997. Intracellular pH regulation. In:, Handbook of Physiology, Cell Physiology. Hoffman JF, Jamieson JD (Eds.) Oxford University Press, New York, pp. 389-440.

154

Queirós AM, Fernandes JA, Faulwetter S, Nunes J, Rastrick SP, Mieszkowska N, Artioli Y, Yool A, Calosi, P, Arvanitidis C, Findlay HS, Barange M, Cheung WWL, Widdicombe S. 2015. Scaling up experimental ocean acidification and warming research: from individuals to the ecosystem. Global Change Biol. 21:130-143.

Quinn TP, Merrill RT, Brannon EL. 1981. Magnetic field detection in sockeye salmon. J Exp Zool A: Ecol Genet Physiol. 217:137-142.

Regan MD, Turko AJ, Heras J, Andersen MK, Lefevre S, Wang T, Bayley M, Brauner CJ, Phuong NT, Nilsson GE. 2016. Ambient CO2, fish behaviour and altered GABAergic neurotransmission: exploring the mechanism of CO2-altered behaviour by taking a hypercapnia dweller down to low CO2 levels. J Exp Biol. 219:109-118.

Réveillac E, Lacoue-Labarthe T, Oberhänsli F, Teyssié JL, Jeffree R, Gattuso JP, Martin S. 2015. Ocean acidification reshapes the otolith-body allometry of growth in juvenile sea bream. J Exp Mar Biol Ecol. 463:87-94.

Roggatz CC, Lorch M, Hardege JD, Benoit DM. 2016. Ocean acidification affects marine chemical communication by changing structure and function of peptide signalling molecules. Global Change Biol. 22:3914-3926.

Rossi T, Nagelkerken I, Pistevos JCA, Connell SD. 2016a. Lost at sea: ocean acidification undermines larval fish orientation via altered hearing and marine soundscape modification. Biol Lett. 12:20150937.

Rossi T, Connell SD, Nagelkerken I. 2016b. Silent oceans: ocean acidification impoverishes natural soundscapes by altering sound production of the world's noisiest marine invertebrate. Proc R Soc B. 283:20153046

Rossi T, Nagelkerken I, Simpson SD, Pistevos JC, Watson SA, Merillet L, Fraser P, Munday PL, Connell SD. 2015. Ocean acidification boosts larval fish development but reduces the window of opportunity for successful settlement. Proc R Soc B. 282:20151954.

Schade FM, Clemmesen C, Wegner KM. 2014. Within- and transgenerational effects of ocean acidification on life history of marine three-spined stickleback (Gasterosteus aculeatus). Mar Biol. 161:1667-1676.

Schmitt BC, Ache BW. 1979. Olfaction: responses of a decapod crustacean are enhanced by flicking. Science. 205:204-206.

155

Shaw EC, Munday PL, McNeil BI. 2013a. The role of CO2 variability and exposure time for biological impacts of ocean acidification. Geophys Res Lett. 40:4685- 4688.

Shaw EC, McNeil BI, Tilbrook B, Matear R, Bates ML. 2013b. Anthropogenic changes to seawater buffer capacity combined with natural reef metabolism induce extreme future coral reef CO2 conditions. Global Change Biol. 19:1632-1641.

Simpson SD, Meekan M, Montgomery J, McCauley R, Jeffs A. 2005. Homeward sound. Science. 308:221-221.

Simpson SD, Munday PL, Wittenrich ML, Manassa R, Dixson DL, Gagliano M, Yan HY. 2011. Ocean acidification erodes crucial auditory behaviour in a marine fish. Biol Lett. rsbl20110293.

Simpson SD, Meekan MG, Larsen NJ, McCauley RD, Jeffs A. 2010. Behavioral plasticity in larval reef fish: orientation is influenced by recent acoustic experiences. Behav Ecol. 21:1098-1105.

Sisneros JA, Tricas TC. 2002. Neuroethology and life history adaptations of the elasmobranch electric sense. J Physiol Paris. 96:379-389.

Spady BL, Watson SA, Chase TJ, Munday PL. 2014. Projected near-future CO2 levels increase activity and alter defensive behaviours in the tropical squid Idiosepius pygmaeus. Biol Open. 3:1063-1070.

Steckbauer A, Ramajo L, Hendriks IE, Fernandez M, Lagos N, Prado L, Duarte CM. 2015. Synergistic effects of hypoxia and increasing CO2 on benthic invertebrates of the central Chilean coast. Front Mar Sci. 2:49.

Thomas ML. 2011. Detection of female mating status using chemical signals and cues. Biol Rev. 86:1-13.

Tierney AJ, Atema T. 1987. Amino acid chemoreception: effects of pH on receptors and stimuli. J Chem Ecol. 14:135–141.

Tolimieri N, Jeffs A, Montgomery J. 2000. Ambient sound as a cue for navigation by the pelagic larvae of reef fishes. Mar Ecol Prog Ser. 207:219-224.

Tomba AM, Keller TA, Moore PA. 2001. Foraging in complex odor landscapes: chemical orientation strategies during stimulation by conflicting chemical cues. J N AM Benthol Soc. 20:211-222.

Urick RJ. 1984. Ambient noise in the sea. Catholic Univ of America, Washington DC.

156

Vermeij MJ, Marhaver KL, Huijbers CM, Nagelkerken I, Simpson SD. 2010. Coral larvae move toward reef sounds. PloS One. 5:e10660.

Walker MM, Dennis TE, Kirschvink JL. 2002. The magnetic sense and its use in long- distance navigation by animals. Curr Opin Neurobiol. 12:735-744.

Watson S-A, Lefevre S, McCormick MI, Domenici P, Nilsson GE, Munday PL. 2013. Marine mollusc predator-escape behaviours altered by near-future carbon dioxide levels. Proc R Soc Lond B. 281:20132377.

Webster NS, Uthicke S, Botté ES, Flores F, Negri AP. 2013. Ocean acidification reduces induction of coral settlement by crustose coralline algae. Global Change Biol. 19:303-315.

Welch MJ, Watson S-A, Welsh JQ, McCormick MI, Munday PL. 2014. Effects of elevated CO2 on fish behavior undiminished by transgenerational acclimation. Nat Clim Change. 4:1086-1089.

Widdicombe S, Spicer JI. 2008. Predicting the impact of ocean acidification on benthic biodiversity: what can animal physiology tell us? J Exp Mar Biol Ecol. 366:187-197.

Wilkens LA, Hofmann MH. 2005. Behavior of animals with passive, low-frequency electrosensory systems. In: Electroreception (pp. 229-263). Springer New York.

Yang XL. 2004. Characterization of receptors for glutamate and GABA in retinal neurons. Prog Neurobiol. 73:127-150.

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Appendix C

PERMISSIONS

C.1 Multiple environmental cues impact habitat choice during nocturnal homing of specialized reef shrimp

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C.2 Impacts of Ocean Acidification on Sensory Function in Marine Organisms

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