Extracellular Ph Changes in Echinoderms As a Consequence of Ocean Acidification

Extracellular pH changes in Echinoderms as a consequence of Ocean Acidification

Esther Stuck

A thesis submitted for the degree of

Master of Science

Department of Marine Science, University of Otago, Dunedin,

New Zealand

June 2014 Abstract .

Atmospheric carbon dioxide levels have recently surpassed 400 ppm, a substantial increase since the 1750's value of 280 ppm. The ocean, an enormous sink of this

CO2, is projected to experience a change in seawater pH of up to 0.77 pH units by the year 2300. Regionally polar latitudes are expected to show the effects of ocean acidification earliest as CaCO3 saturation levels are already low due to cold waters.

This research aimed to develop and test a technique that allowed the measurement of extracellular pH (pHe) in developing echinoderm larvae when exposed to pH values predicted for the future. Morphological changes were also recorded in parallel with pHe. To investigate if responses varied with latitude, I looked at the effect of lowered seawater pH on the pHe of five echinoderm species: the tropical sand dollar Arachnoides placenta, the temperate summer spawning urchin Evechinus chloroticus, and temperate winter spawning Pseudechinus huttoni, the polar sea urchin Sterechinus neumayeri and the polar sea star Odontaster validus. Different developmental stages were used (blastula, gastrula, pluteus and bipinnaria) to investigate how pHe regulation changed during development. Body regions of the developing larvae were also distinguished (gut, oesophagus and arm) to identify if pHe regulation differed throughout the pluteus and bipinnaria larvae. To determine the affects of ocean acidification, pH levels of ambient pH, pH 7.8, pH 7.6, and pH 7.4 were used for the tropical and temperate experiments and ambient pH, pH 7.7, pH 7.5 and pH 7.2 for the polar experiments.

The technique to measure extracellular pH was developed using the nontoxic fluorescent probe 8-hydroxypyrene-1,3,6-trisulfonic acid (HPTS). After incubation of larvae in 30 µM HPTS, dual photographs were taken using the probes excitation optimums of 405 nm and 460 with a single emission optimum at 520 nm to produce a false colour pH map of larval pHe in MATLAB.

The temperate sea urchin E. chloroticus showed the least tolerance to lowered seawater pH as pHe was significantly decreased in the gut (pHe 7.91 to pHe 6.5), oesophagus (pHe 6.59 to pHe 5.97) and arms (pHe 6.65 to pHe 6.17) when raised under pH 7.6. Sterechinus neumayeri blastula did not show an effect to lowered pH but the pluteus larvae had significantly decreased pHe of up to 0.44 units (gut pHe) in all regions in treatment pH 7.2. The temperate P. huttoni was the most tolerant to a lowering seawater pH with only the gut region responding with a significantly lowered pHe (pHe 6.67 to pHe 6.64) in treatment pH 7.4. Overall polar species had the lowest pHe in all regions which decreased as seawater pH decreased. A 0.4 pH unit drop in pH (pH 7.6-7.8) resulted in significant abnormality of the early cell stage and size reductions of the pluteus in all species and sea star bipinnaria.

The effects of ocean acidification on the ability of echinoderms to regulate their extracellular pH are species-specific. Several parameters, such as taxonomic

i differences, physiology, genetic makeup and the population’s evolutionary history may contribute to this variability. This study highlights that although many species may be negatively affected by ocean acidification, the ability for polar species to regulate pHe is surprisingly effective; they are only affected when pH decrease is extreme. Further research is needed, especially looking at multiple stresses, as ocean acidification will not happen on its own but in conjunction with ocean warming, increases in UV-B irradiance, overfishing and pollution. The technique developed in this study will allow insight into how internal systems of many species cope with climates associated changes.

Acknowledgements .

I would like to express my deep gratitude to my research supervisor, Dr Miles Lamare for the opportunity to work on a great project and for his guidance and support throughout this journey. Thank you to my co-supervisor Abby Smith for her additional support, enthusiastic encouragement and useful critiques of this research work.

My grateful thanks are also extended to Kim Currie and the Belgium crew on the ice for their help in analysing water chemistry data. Thanks to Suse and Cerys who helped design the MATLAB coding. Thank you to Jean for her fantastic starfish drawing. A big thank you to the technicians at the Portobello Marine Science laboratory especially to Albie, Reuben and Dave for going out of their way to help me on many occasions.

I am thankful for the opportunity to have gone to the Australian Institute of Marine Science (AIMS) to carry out research and a special thanks Sven Uthicke for his great hospitality and laughs. Thank you to Maria Byrne for her expertise and endless energy in Antarctica. Thanks to Paul Meredith for being a superb skipper and Mattie B and Rory for being around to help collect urchins for me.

A big thank you to the rest of the staff at NZMSS who were always smiling and willing to help in any way, especially Steve Cutler for giving me the opportunity to gain work experience while doing my thesis. In terms of funding I would like to express my great appreciation to the Otago Branch of the Federation of Graduate Women for the Brenda Shore award and OMV Ltd. for their financial support towards my research costs.

Thank you to Buzzy and Olga for being great role models and travel companions. A big thank you to all my friends and classmates who made the last two years memorable, you guys rock! Finally, I wish to thank my parents for their love and support throughout my time here in Dunedin.

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Table of Contents . .

Abstract i

Acknowledgements iii

Tables of Contents iv

List of Tables vii

List of Figures x

List of Appendices xviii

Chapter 1: General Introduction 1

1.1 Ocean acidification 2

1.2 Saturation of calcite and aragonite 6

1.3 Marine life in an acidifying ocean 10

1.4 Acid base regulation 19

1.5 Aims and objectives 25

Chapter 2: Measuring extracellular pH (pHe) 27

2.1 Introduction 27

2.2 Methods 30

2.2.1 Fluorometry 30

2.2.2 Measuring the difference in emission/absorption 30

2.2.3 Larval photo analysis 33

Chapter 3: Response of Extracellular pH (pHe) to Reduced Seawater pH in Larvae of Five Echinoderm Species 37

3.1 Introduction 37

3.1.1 General latitudinal trends in seawater carbonate chemistry 37

3.2 Aims 42

3.3 Study species: Evechinus chloroticus, Pseudechinus huttoni, Arachnoides placenta, Sterechinus neumayeri, and Odontaster validus 42

3.4 Materials and methods 47

3.4.1 Animal collections 47

3.4.2 Spawning & culturing 49

3.4.3 Adjustment and monitoring of seawater parameters 51

3.4.4 Morphometrics 55

3.4.5 Extracellular pH measurements 56

3.4.6 Statistical analysis 58

3.5 Results 59

3.5.1 Adjustment and monitoring of seawater parameters 59

3.5.2 Morphological observations of early cell development 62

3.5.3 Morphological observations of Larvae - gastrula, pluteus and sea star bipinnaria developmental stages 65

3.5.4 Extracellular pH (pHe) of Larvae - gastrula, pluteus and seastar bipinnaria developmental stages 69

3.6 Discussion 87

3.6.1 Adjustment of seawater pH and ambient pH 87

3.6.2 Extracellular pH measurements in echinoderms 88

3.6.3 Morphological observations with lowered pH 93

3.6.4 Physiological responses 98

3.6.5 Latitudinal differences 105

Chapter 4: General discussion and conclusion 110

4.1 Summary of results 110

4.2 Ecological impacts 112

4.3 Climate change and multiple stressors 114

4.4 Future research 115

4.5 Conclusions 117

References 118

Appendices 132

List of Tables .

1.1 Predicted increases in atmospheric CO2 with ocean surface temperature, pH change and aragonite saturation. 7

1.2 Images of the physical effects of increased seawater CO2 on a variety of marine organisms, including species of a) mollusc, pteropod, foraminifera and b) echinoderms 15

3.1. Details on the collection location, experimental conditions and reproductive period of the five study species: Evechinus chloroticus, Pseudechinus huttoni, Arachnoides placenta, Sterechinus neumayeri and Odontaster validus 45

3.2. Temperature (°C), salinity (PSU), pH (NBS scale), total alkalinity (µmol kg-1 1 soln- ), partial pressure of CO2 (ρCO2), CaCO3 saturation value for calcite (ΩC)

and aragonite (ΩA) for ambient and treated seawater. (a) Evechinus chloroticus (c) Pseudechinus huttoni (d) Arachnoides placenta (d) Odontaster validus (e) Sterechinus neumayeri 61

3.3 Students t-test to identify significant asymmetrical differences between the post oral arm 1 and post oral arm 2 on the pluteus from a) E. chloroticus early b) A. placenta c) P. huttoni , d) S. neumayeri . n=10 68

3.4. One-way ANOVA for Evechinus chloroticus pluteus to identify significant differences in pHe among pH treatments. Body components tested are gut (a), gut wall (b), oesophagus (c) and arms (d). Data were log transformed and a Bartlett test confirmed homogeneity of variance. Heterogeneous variances were compared using Welch ANOVA testing for equal means, allowing for unequal variances 71

3.5. One-way ANOVA for Pseudechinus huttoni pluteus to identify significant differences in pHe among pH treatments. Body components tested are gut (a), oesophagus (b) and arms (c). Data were log transformed and a Bartlett test

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confirmed homogeneity of variance. Heterogeneous variances were compared using Welch ANOVA testing for equal means, allowing for unequal variances 75

3.6. One-way ANOVA for Arachnoides placenta pluteus to identify significant differences in pHe among pH treatments. Body components tested were gut (a), oesophagus (b) and arms (c). Data were log transformed and a Bartlett test confirmed homogeneity of variance. Heterogeneous variances were compared using Welch ANOVA testing for equal means, allowing for unequal variances 77

3.7. One-way ANOVA for Sterechinus neumayeri blastula to identify significant differences in pHe among pH treatments. Body components tested were the blastocoel (a) and blastoderm (b). Data were log transformed and a Bartlett test confirmed homogeneity of variance. Heterogeneous variances were compared using Welch ANOVA testing for equal means, allowing for unequal variances 79

3.8. One-way ANOVA for Sterechinus neumayeri pluteus to identify significant differences in pHe among pH treatments. Body components tested were the gut (a), oesophagus (b) and arms (c). Data were log transformed and a Bartlett test confirmed homogeneity of variance. Heterogeneous variances were compared using Welch ANOVA testing for equal means, allowing for unequal variances 81

3.9. One-way ANOVA for Odontaster validus gastrula to identify significant differences in pHe among pH treatments. Body components tested were the oral hood (a) and gut (b). Data were log transformed and a Bartlett test confirmed homogeneity of variance. Heterogeneous variances were compared using Welch ANOVA testing for equal means, allowing for unequal variances 81

3.10. One-way ANOVA for Odontaster validus 43 day old bipinnaria larvae to identify significant differences in pHe among pH treatments. Body components tested were the gut (a), oral hood (b) and oesophagus (c). Data were log transformed and a Bartlett test confirmed homogeneity of variance. Heterogeneous variances were compared using Welch ANOVA testing for equal means, allowing for unequal variances 85

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3.11. Measurements of extracellular pH from four adult sea urchins, one pluteus sea urchins and one adult sea star, when reared under ambient seawater conditions 90

3.12. pHe values of the pluteus/bipinnaria regions (gut, oesophagus and arms/oral hood) of: a) A. placenta (n = 10), b) E. chloroticus (n=8), c) P. huttoni (n=10), d) Sterechinus neumayeri (n=10) and e) Odontaster validus when exposed to lowered seawater pH. Number in brackets indicated standard error 92

3.13 Examples of the effects of reduced seawater pH on the different early developmental stages and survival of a range of marine calcifiers from tropical, temperate and polar regions 96

List of Figures .

1.1. Hourly, daily and monthly averages of carbon dioxide concentrations at Mauna

Loa, Hawaii. This is the first month in which every day had a CO2 concentration above 400ppm (Thompson, 2014) 3

1.2. The distribution of inorganic carbon species as a function of seawater pH. Present-day seawater values are marked by the thick vertical red line (modified from Erez et al., 2011 p.6) 4

1.3. Predicted ocean CO2(aq) increase along with seawater pH decrease (Feely et al., 2006). 6

1.4. Distribution of (A) aragonite and (B) calcite saturation depth (Ω = 1) in the global oceans. The level at which aragonite and calcite are in thermodynamic equilibrium is known as the saturation depth. When the degree of saturation, Ω, is greater than 1, seawater is supersaturated with aragonite and calcite; conversely, seawater is under saturated with respect to these minerals when Ω < 1. This depth is significantly shallower for aragonite than for calcite, because aragonite is more soluble in seawater than calcite (Feely et al., 2004, p.2). 8

1.5. Estimated aragonite saturation states of the surface ocean for the years 1765, 1995, 2040, and 2100, this is an example for shallow and deep corals under

predicted CO2 emission scenarios (Kleypas et al., 2006 p.17) 9

1.6. Absorbed CO2 in seawater (H2O) forms carbonic acid (H2CO3), lowering the water's pH level and making it more acidic. This raises the hydrogen ion concentration in the water, and limits organisms' access to carbonate ions, which are needed to form hard parts (modified from Kleypas et al., 2006, p.15; UKOA 2013) 11

1.7. Life cycle of the seastar Patiriella regularis. Different life stages will have differing sensitivities to climate change stressors (modified from Byrne 2011, p.3). 13

1.8. Evechinus chloroticus larvae under normal light microscope (left) and Arachnoides placenta larvae under polarised light, showing larval skeleton (right). Scale bars = 100µm. 14

1.9. Seasonal estimates of pH and CO32- for the Southern Ocean. (Top) Winter and summer distributions of pH. (Middle) Winter and summer distributions of CO32- (McNeil and Matear, 2008, p.2). 18

1.10. A cross section of a sea urchin spicual during biomineralization. 1, various trans-membrane transporters for HCO3- and calcium from surrounding; 4,

action of carbonic anhydrase (CA), an enzyme that converts CO2 in the cell; 5,certain spicule matrix proteins (SM) are involved in directing biomineralization in the extracellular space between the PMC and the growing skeleton (Hofmann et al., 2008, p.4). 21

1.11. Schematic model summarizing the interplay of calcification and pH regulation. pH homeostasis is maintained by ion transporters, which either directly or indirectly depend on the consumption of energy. Increased energetic costs due to decreased proton gradients lead to a shift in the larvae’s energy budget, which decreases scope for growth and may also translate into juvenile impaired fitness. ST, stomach; (Stumpp et al., 2004, p.4). 24

2.1. pH dependence of the absorption spectrum of HPTS . Maximum absorbance is shaded red ~ 405nm(λ1 exc) , ~460nm(λ2 exc ) (modified from Wolfbeis et at., 1983). 30

2.2. Plot of HPTS fluorescence intensity at 520 nm as a function of the pH-value at various excitation wavelengths. When pH is alkaline the HPTS solution is bright green/yellow (right), this disappears gradually as the pH is lowered (left) (modified from Wolfbeis et al., 1983). 31

2.3. An example of fluorescent images and ratios produced by 3 pH calibrations from the addition of 1M of HCL and NaOH to 30µM HPTS seawater solution. Excited at wavelengths 460nm(λ2exc) and 405nm(λ1exc), and photographed at 520nm using a set exposure of 2 seconds. 32

2.4. Example of calibration curve for 30µM HPTS. Equation and R2 value of the polynomial. Note only eight points shown as readings above pH 8.5 become saturated at a ratio of 1 and below 5 a ratio of 0 (no florescence) so readings at these values were removed 33

2.5. MATLAB code used to produce pH profile of larvae. 1) MATLAB reading images 405nm (blue) and 460nm (green) 2) Equations for each species, from standard curves 3) pH profile with limits between pH 6-9. 4) Allows manual isolation of regions and calculates the average pH within that region 35

2.6. An example of a 1 day old A. placenta larvae (from top left), normal, 405nm, 406nm, and the MATLAB generated pH colour map. The red dashed lines indicated where average pH measurements were made on at the pluteus stage, this included the gut(G), larval arms (A), mouth (M), and in some cases the gut wall(GW) if it was defined. 36

2- 3.1. Increasing atmospheric CO2, decreasing surface ocean pH and [CO3 ] over time and latitude predicted for the year 2100 (a) lead to reductions in surface ocean 2- pH (b) and surface ocean [CO3 ]. Results are given as global averages from

1994 data and the preindustrial (‘Preind.’) and include two future CO2 scenarios (IS92a ‘continually increasing’ scenario of 788 ppm in the year 2100 and S650 ‘stabilization’ scenario of 563 p.p.m.v. in the year 2100). The thin 2- dashed lines indicate the [CO3 ] for sea water in equilibrium with aragonite and calcite are nearly flat, revealing weak temperature sensitivity (Modified from Orr et al 2005 p.682). 38

3.2. Photographs of adult echinoderms used in this study: A. Arachnoides placenta B. Evechinus chloroticus, C. Pseudechinus huttoni, D. Sterechinus neumayeri,

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E. Odontaster validus. Photographs were taken by author. Scale bars represent 10mm 46

3.3. Maps showing location of (a) Doubtful Sound, Fiordland, New Zealand, where E. chloroticus and P. huttoni were collected (Worldatlas.com, 2013). (b) Palleranda Beach, where tropical sand dollar A. placenta was collected (About.com, 2013) and (c) Cape Evans, Antarctica, (Tremblay, 2013) where S. neumayeri and O. validus were collected. 48

3.4. Image of spawning female S. neumayeri (left) and male P. huttoni (right) after being injected with KCL to induce spawning. Images taken by author 50

3.5. 50ml falcon tube used to rear larvae in flow through seawater systems. Four, 5x0.5cm cuts were made around tube, covered by 100µm mesh to keep larvae in but still allow water to flow through. Scale bar represents 1cm. 50

3.6. pH controlled flow-through larval rearing system for New Zealand experiments with 1, 2 and 3 representing the replicate tanks...... 52

3.7. Photos of flow through systems for each species: a. E. chloroticus/P. huttoni, b. A placenta, and c. S. neumayeri 54

3.8. Morphometric measurements of a) sea urchin larvae, using Evechinus chloroticus as an example and b) Odontaster validus gastrula and bipinnaria. Measurements made are total length (TL), total width (TW), and post-oral skeletal arms 1 and 2 (R1, R2) on pluteus larvae. Pluteus larvae were also viewed under normal and polarised light to highlight the larval skeleton. Scale bar represents 100 µm.. 56

3.9 Images of larvae after being incubated in HPTS, washed, excited at wavelengths 405 nm/495 nm and photographed at emission 520 nm (Odontaster validus as an example). Scale bar represents 100 µm.. 57

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3.10. Early embryological development of E. chloroticus, P. huttoni, A. placenta, S. neumayeri and O. validus when raised under four seawater pH treatments. Scale bars represent 50µm 63

3.11. Percentage of a) Abnormal early egg development, b) Abnormal 2-cell embryonic stage c) Abnormal 4-cell embryonic stage and d) Abnormal blastula, of five echinoderm species when larvae were grown in either ambient pH seawater, or lowered pH seawater pH. Different letters indicates significant differences between pH treatments (P<0.05) (N = 30) 64

3.12. Length of morphometric variables in larvae of five species grown in either ambient pH seawater, or lowered pH seawater a) Evechinus chloroticus, b) Arachnoides placenta, c) Pseudechinus huttoni, d, e) Sterechinus neumayeri and f, g) Odontaster validus. While similar in developmental stage (4-armed pluteus or gastrula), larvae varied in age. Columns having different lower case indicates significant differences among pH treatments (P<0.05) (N = 30) 67

3.13. Mean length of post oral arms 1 and 2 for a) E. chloroticus b) A. placenta c) P. huttoni and d) S. neumayeri under ambient and three reduced seawater pH levels. n =10. Error bars are ± standard error.. 68

3.14. Images of 12 day old Evechinus chloroticus larvae and resulting false colour pH map produced in MATLAB when raised under varying seawater pH. Scale bar represents100µm.. 71

3.15. Mean3extracellular pH in Evechinus chloroticus under ambient pH and three reduced seawater pH levels for the gut (a), gut wall (b), oesophagus (c) and post oral arms (d). Error bars are ± standard error. Significant differences among pH treatments are indicated by columns not sharing the same letter. n=8.. 73

3.16. Images of 12 day old Pseudechinus huttoni larvae and resulting false colour pH map produced in MATLAB when raised under ambient pH and three reduced seawater pH levels. Scale bar represents 100µm 74

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3.17. Mean extracellular pH in Pseudechinus huttoni under ambient pH and three reduced seawater pH levels for the gut (a), oesophagus (b) and arms (c). Error bars are ± standard error. Significant differences among pH treatments are indicated by columns not sharing the same letter. n=10 76

3.18. Images of four day old Arachnoides placenta larvae and resulting false colour pH map produced in MATLAB when raised under ambient pH and three lowered seawater pH levels. Scale bar represents100µm 77

3.19. Mean extracellular pH in Arachnoides placenta under ambient pH and three reduced seawater pH levels for gut (a), oesophagus (b) and arms (c). Error bars are ± standard error. Significant differences among pH treatments are indicated by columns not sharing the same letter. n=9 78

3.20 Images of seven day old Sterechinus neumayeri larvae and resulting false colour pH map produced in MATLAB when raised under varying seawater pH. Components tested were the blastocoel (a) and blastoderm (b). Scale bar represents 50µm 79

3.21. Mean extracellular pH in Sterechinus neumayeri blastula under ambient pH and three reduced seawater pH levels for the blastocoel (a) and blastoderm (b). Error bars are ± standard error. Significant differences among pH treatments are indicated by columns not sharing the same letter. n=10 80

3.22. Images of 4 week old Sterechinus neumayeri larvae and resulting false colour pH map produced in MATLAB when raised under varying seawater pH. Scale bar represents 50µm 80

3.23. Mean extracellular pH in Sterechinus neumayeri under ambient pH and three reduced seawater pH levels for the gut (a), oesophagus (b) and arms (c). Error bars are ± standard error. Significant differences among pH treatments are indicated by columns not sharing the same letter. n=10 82

3.24. Images of 15 day old Odontaster validus larvae and resulting false colour pH map produced in MATLAB when raised under varying seawater pH. Scale bar represents 100µm 83

3.25. Mean extracellular pH in Odontaster validus gastrula under ambient and reduced seawater pH levels for the oral hood (a) and gut (b). Error bars are ± standard error. Significant differences among pH treatments are indicated by columns not sharing the same letter. n=10 84

3.26. Images of 43 day old Odontaster validus larvae and resulting false colour pH map produced in MATLAB when raised under varying seawater pH. Scale bar represents 100µm 85

3.27. Mean extracellular pH in 43 day old Odontaster validus bipinnaria larvae under ambient and reduced seawater pH levels for the gut (a), oral hood (b) and oesophagus (c). Error bars are ± standard error. Significant differences among pH treatments are indicated by columns not sharing the same letter. n=10 86

3.28. a) Relationship between seawater and the primary body cavity pH (NBS scale) of Strongylocentrotus droebachiensis pluteus during acute and chronic environmental hypercapnia. Bars represent ±SD; n = 4–5. (Stumpp et al. 2012 p.18193) b) Coelomic fluid pH (pH CF) of Paracentrotus lividus according to the aquarium seawater pH (pHSW) (Catarino et al., 2012 p 2348) c) Changes in pHe in the coelomic fluid after exposure to hypoxia in Psammechinus miliaris, data are means + SD (n= 6 ) ( = coelornic fluid; = surrounding water) (Spicer et al. 1995 p 73) 91

3.29. Ion regulation in invertebrate larvae during hypercapnia. When acidity increases, extracellular protons inhibit (-) the Na+/H+ exchanger. As a + + - - consequence, the Na /H /Cl /HCO3 exchanger becomes increasingly involved in pHi regulation while reducing in Na+/K+-ATPase activity (modified from Portner et al., 1998 p1806) 102

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3.30. Reduced growth of the postoral arm in echinoplutei in response to changes in calcite and aragonite saturation. The solid line is the regression line and the shaded area is 95% CI. Different colours represent geographical regions. (Byrne et al., 2013c). 106

3.32. Changes in gut pHe in response to altered seawater pH. Species were pooled into either tropical (A. placenta), temperate (E. chloroticus and P. huttoni) and polar (S. neumayeri and O. validus) latitudinal regions 107

3.33. Latitude Vs mean pHe of the gut, oesophagus and arms regions in our five study species. Error bars are ± 1SE 108

4.1. Ocean acidification can directly (delays in growth/calcification) or indirectly (longer in the plankton increases predation) affect many physiological processes with consequences on fitness and survival (Modified from Dupont and Thorndyke, 2008 p3131) 113

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List of Appendices .

Appendix 1. The effects lowered seawater has on early cell development. pH ambient: predominantly cells are spherical shape and blastomeres are even in size. pH 7.8/7.7: most species cells are still spherical in shape, with regards to slightly abnormal cells in P. huttoni and a slightly unsmooth blastoderm in A. placenta blastula. pH 7.6/7.5: delayed cell development in E. chloroticus, irregular embryos for P. huttoni and S. neumayeri. pH 7.4/7.2: Delayed cell development for E. chloroticus, delayed and abnormal cell development for P. huttoni, abnormal blastoderm development of A. placenta, irregular shaped cells of S. neumayeri, O. validus has irregular blastula development with disrupted cell organisation around the blastoderm. 132

Appendix 2. Ratio of λ2/λ1 (405/460) after emission at 520 nm (x-axis) as a function of seawater pH (y-axis) using the pH sensitive dye HPTS. Black dots indicate pH reading at specific ratios. A calibration curve was fitted using a non linear polynomial and a non linear regression equation was obtained for each experiment; a. E. chloroticus, b. P. huttoni, c. A. placenta, d. O. validus e. S. neumayeri.. Standard errors are smaller than graph points. 133

Appendix 3. One-way ANOVA on the abnormality of early cell development for a) E. chloroticus early b) A. placenta c) P. huttoni , d) S. neumayeri and d) O. to identify significant differences among pH treatments. Embryos were grown in ambient and three lowered seawater pHs. 135

Appendix 4. One-way ANOVA on the length of morphometric variables for a) E. chloroticus early b) A. placenta c) P. huttoni , d) S. neumayeri and d) O. validus to identify significant differences among pH treatments. While similar in developmental stage (4-armed pluteus or gastrula), larvae varied in age. 137

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Chapter 1 General introduction

Due to anthropogenic emissions of CO2, a substantial increase in atmospheric CO2 concentration from the 1750's ρCO2 of 280 ppm (Bernstein et al., 2008) first exceeded 400 ppm on May 9th 2013 (Kunzig 2013), with April 2014 becoming the first month with an average CO2 concentration above 400 ppm (Thompson, 2014). Since the ocean is an important sink of the CO2 humans produce, it has experienced fundamental changes in the chemistry of seawater leading to an increase in the concentration of CO2 in surface waters (Portner, 2008, Raven et al., 2005). This increase in CO2 leads to an increase in hydrogen ions (H+), causing seawater to become less alkaline and as a result, a decrease in the carbonate saturation state of the Earth’s oceans (Orr et al., 2- 2005); a process known as ocean acidification. Carbonate ions (CO3 ) are crucially important in the production of skeletons and shells for many marine organisms (Stumpp et al., 2012). The increase in H+ reacting with carbonate ions, overall causes a decrease 2- in CO3 ion concentrations. With continued ocean acidification many parts of the ocean are projected to become undersaturated with respect to calcium carbonate minerals (Doney et al., 2009), leaving marine calcifiers with less material to build their skeletons from (Caldeira and Wickett, 2003, Wolf-Gladrow et al., 1999).

Echinoderms are among the heavily calcified marine organisms that may be affected by ocean acidification (Brennand et al., 2010, Clark et al., 2009, Dupont et al., 2008, Gonzalez-Bernat et al., 2013a, Gonzalez-Bernat et al., 2013b, Kurihara, 2008). Vital feeding and skeletal structures of echinoderms are calcified (Bentov et al., 2009, Pelletier, 2010). Larval stages are particularly vulnerable to acidification as they are small and tend to produce a soluble polymorph of carbonate (Politi et al., 2004). Polar larvae which live in a naturally higher CO2 environment and extremely cold waters

1 have very slow metabolisms which may be more or less sensitive to acidification and act as a bellwether for other echinoderm species (Bednarsek et al., 2012). As well as the effects on calcification, ocean acidification may influence extracellular acid–base regulation, which plays an important role in calcification rates and other physiological processes of many marine calcifiers, such as ion pumping (Brownlee 2009).

This Master's thesis investigates the capacity of echinoderm larvae to regulate extracellular pH under increased seawater CO2. In order to establish variation within the group, larval stages from five ecologically important and latitudinal differing echinoderm species are investigated: one tropical: Arachnoides placenta, two temperate: Evechinus chloroticus and Pseudechinus huttoni and two polar: Sterechinus neumayeri and Odontaster validus.

1.1 Ocean acidification

Greenhouse gases are drivers of climate change; global emissions due to human activities caused an atmospheric carbon dioxide (CO2) increase of 80% since 1750 which is expected to rise another 40% by 2100 (Bernstein et al., 2008). The global atmospheric concentration of ρCO2 increased from a pre-industrial (~1750’s) value of

280ppm to 379ppm in 2005 (Bernstein et al., 2008). The present CO2 concentration, surpassing a monthly average above 400ppm in April 2014 (Figure 1) (Thompson, 2014), is the highest in the atmosphere for more than 4.5 million years, (since the Pliocene) (Bartoli et al., 2011) and possibly in the last 20 million years (Tripati et al., 2009). This increase is expected to continue and will lead to significant global temperature increases by the end of the century (Cicerone et al., 2004).

Figure 1.1 Hourly, daily and monthly averages of carbon dioxide concentrations at

Mauna Loa, Hawaii. This is the first month in which every day had a CO2 concentration above 400ppm (Thompson, 2014, Fig 2).

The ocean covers 70% of Earth’s surface; due to its large volume it is an enormous sink of anthropogenic carbon, absorbing approximately one third of the CO2 that has been produced since 1750 (Cicerone et al., 2004). Dissolution of CO2 however has led to the ocean becoming less alkaline with an average global decrease in pH of 0.1 units (recorded in 1996) since pre-industrial times, equating to a 30% increase in hydrogen ions (Bernstein et al., 2008, Henry, 1803, Raven et al., 2005). Sea surface temperatures have also been higher during the past three decades since at least the 1880’s (NOAA, 2012a). At present, the oceans take up about 2 of the 6 Gt of carbon per annum from human activity (Portner, 2008), twice as much as the terrestrial biosphere. Nevertheless the ability of the ocean to take up CO2 decreases with increasing atmospheric CO2, following in accordance with Henry’s law (Henry, 1803). Therefore, increasing atmospheric CO2 levels directly increases the concentration of CO2 in surface waters

(Portner, 2008, Raven et al., 2005). Additionally CO2, which is relatively unreactive in air, becomes much more reactive when absorbed by the ocean, resulting in an increase in the concentration of hydrogen ions and various negatively charged forms of dissolved carbon (Raven et al., 2005), highlighted in the following equations:

CO2(g) ↔ CO2(aq) (1)

CO2 + H2O ↔ H2CO3 (2) - + H2CO3 ↔ HCO3 + H (3)

- 2- + HCO3 ↔ CO3 + H (4)

The overall effect of CO2 dissolving in seawater is an increased concentration of hydrogen ions (H+) and the production of four main inorganic forms known collectively as dissolved inorganic carbon (DIC): (1) CO2 (CO2(aq)), which is hydrated to form (2) carbonic acid, (H2CO3, as aqueous CO2 can be in either form). This weak acid readily releases the hydrogen ions to form the other types of dissolved inorganic carbon such as - 2− (3) bicarbonate (HCO3 ) and (4) carbonate ions (CO3 ) (Erez et al., 2011, Raven et al., 2005, Seinfeld and Pandis, 2012). Each of these carbon forms varies in abundance due to seawater temperature, salinity and pressure. Therefore, under current ocean − conditions, HCO3 is the most abundant (91%) form of CO2 dissolved in seawater, with 2- CO2(aq) making only 1% and CO3 , 8% (Figure 1.2) (Raven et al., 2005). As oceans become more acidic the graph moves to the left and changes in CO2(aq) increase 2− dramatically while CO3 on the other hand is largely decreased.

Figure 1.2 The distribution of inorganic carbon species as a function of seawater pH.

Present-day seawater values are marked by the thick vertical red line (modified from Erez et al., 2011 p.6) .

The proportions of the three types of DIC reflect the pH of seawater, which acts as a natural buffer when CO2 (acid) is added. This buffer works by additional hydrogen ions 2- 3- + reacting with CO3 , converting them to HCO , reducing H (acidity) ion concentration and therefore acidity, such that the pH change is much less then would be expected. The capacity of this buffer to restrict pH changes decreases as increased amounts of 2- CO2 are absorbed by the oceans, due to the reduction of CO3 which are required for the buffer (Raven et al., 2005). All three forms of dissolved CO2 are important for biological activities of marine organisms such as photosynthesis, the production of complex organic carbon molecules from sunlight, and calcification. When these organisms die or are consumed, most of the carbon stays in the surface waters or is released back into the atmosphere. Some organic material can, however, fall into deep water, in addition to the transport of the CO2 from ocean surfaces by ocean currents (Portner, 2008). This process is referred to as the ‘biological pump’. By transferring carbon from surface waters to greater depths, the pump increases the capacity for the oceans to act as a sink for atmospheric CO2 (Raven et al., 2005). Any changes in the strength of this pump would have significant consequences on the amount of carbon being transported to the deep ocean environments and therefore removed from the atmosphere (Raven et at., 2005).

Since the year 2000, atmospheric CO2 levels have been increasing approximately 100- fold (Levinson and Lawrimore, 2008), 2.33 ppm/year recorded in 2012 (NOAA, 2012b) compared with the end of the last ice age where levels rose by about 80 ppm over a 6000 year period (Bernstein et al., 2008). With the continued use of fossil fuels, atmospheric CO2 concentrations are expected to rise from the current ρCO2 (400 ppm) (Kunzig, 2013) to ~750 ppm (Bernstein et al., 2008, Houghton et al., 2001) or ~1000 ppm by 2100 (Karl et al., 2009). This concentration is expected to increase further to ~1500 ppm between 2100 and 2200 (Raven et al., 2005). This increase will lead to a pH reduction in the upper ocean layers of 0.3 to 0.5 units (Caldeira and Wickett, 2003, Raven et al., 2005), resulting in a pH of 7.8 around the year 2100, representing an ocean that is 320% more acidic than it was in pre-industrial times (Figure 1.3) 2- (Guinotte and Fabry, 2008). With seawater pH dropping to between 7.8 and 7.9, CO3 concentrations would decrease by at least 50% as H+ ions in the seawater reacted with them. Finally, acidification of the surface water by up to 0.77 pH units (ph <7.5) is expected if values of atmospheric CO2 achieve levels of 1900 ppm by 2300 (Caldeira

5 and Wickett, 2003, Wolf-Gladrow et al., 1999). Under such conditions, marine calcifiers would have substantially less material to maintain their shells and skeletons.

Figure 1.3 Predicted ocean CO2(aq) increase along with seawater pH decrease (Feely et al., 2006).

1.2 Saturation of calcite and aragonite

The degree of saturation of seawater with respect to a mineral (aragonite or calcite) is 2+ 2- known as Ω. It is the ion product of the concentrations of calcium (Ca ) and CO3 , at the in situ temperature, salinity, and pressure, divided by the stoichiometric solubility product (K*sp) for those conditions (Feely et al., 2004). It is a measure of the potential for the mineral to precipitate or dissolve. When Ω falls below 1, the saturation horizon, dissolution outweighs production. Calcium carbonate mineral phases are defined by:

2+ 2– 2+ 2– Ωarag = [Ca ][CO3 ] or Ωcal = [Ca ][CO3 ]

K*sparag K*spcal

2+ 2- Here, K*sp is the concentration of Ca and CO3 when the mineral is at equilibrium (not forming nor dissolving) (Atkinson and Cuet, 2008, Feely et al., 2009, Feely et al., 2- 2004). Figure 1.2 shows how decreasing ocean pH leads to a decrease in CO3 concentration causing a reduction in the saturation state of CaCO3, making dissolution more likely (Raven et al., 2005, Wolf-Gladrow et al., 1999). CaCO3 occurs in two

polymorphs, calcite and the more soluble form aragonite, both important in calcifying organisms (Doney et al., 2009, Orr et al., 2005). Ocean surfaces are generally saturated with respect to aragonite, however this level of saturation changes with temperature, pressure, and depth. As biogenic calcium carbonate particles fall through the water column they begin to dissolve below the depth at which Ω = 1 (Woosley et al., 2012).

In the Southern ocean the lysocline (depth in the ocean where the solubility of calcium carbonate increases dramatically) generally occurs around 1,000m, but as of 2004 this lysocline had shoaled by 40-200m as a direct consequence of the uptake of

anthropogenic CO2 (Feely et al., 2004, Orr et al., 2005). Even more recently, studies in

the Southern Ocean observed the aragonite saturation (ΩA) point was reached at 200m depth, through a combination of ocean acidification and natural upwelling (Bednaršek

et al., 2012). Model simulations predict that in 2100 the Southern Ocean’s average ΩA will decrease to 60m, and that the entire water column in the Weddell Sea will become

undersaturated (Orr et al., 2005). Increasing CO2 levels and the resulting lower pH of

seawater decreases the saturation state of CaCO3 and raises the saturation horizons of both aragonite and calcite closer to the surface (Doney et al., 2009). This decrease in saturation state is believed to be one of the main factors leading to decreased calcification in marine organisms (Marubini et al., 2008). Organisms that produce aragonite may be particularly vulnerable to changes in ocean acidity relative to those

that produce calcite (Table 1.1).

Table 1.1. Predicted increases in atmospheric CO with ocean surface temperature, pH 2 change and aragonite saturation .

Atmospheric CO2 Time Period Average ocean Ocean Aragonite (ppm) (Years) surface temp (°C) pH saturation

180 18000BP 12.3 8.32 4.26

280 1880 15.9 8.16 3.44

393 2010 16.5 8.05 2.9

450 2040-2050 17.6 7.95 2.5

560 2050-2070 18.3 7.91 2.29

840 2100 20 7.76 1.81

1900 2300 >23 7.43 0.90

(Collated from Caldeira and Wickett 2003; Kleypas et al., 2006; CO2 speciation calculator) BP = before present 7

Increasing oceanic CO2 uptake will drive waters to a point where aragonite will start to dissolve (Table 1.1). Antarctic waters are expected to experience this first (McNeil et al., 2010), due to the naturally lower calcium carbonate (CaCO3) concentration in the

Southern Ocean. CO2 absorption is enhanced by increasing solubility at low water temperatures, whereas warming favours CO2 release (Portner, 2008). In the Indian Ocean, saturation depths have become shallower increasingly north of 30°S, equating to aragonite saturation depths in the Arabian Sea and Bay of Bengal 100-200m shallower than in pre-industrial times (Feely et al., 2004). In the Pacific, the upward migration of the aragonite saturation horizon is 30-80m south of 38°S and 30-100 m north of 3°N. The calcite saturation horizon has also shoaled by about 40-100 m north of 20°N in the North Pacific (Figure 1.4). The dissolution of CaCO3 particles is likely to increase as the waters become increasingly under-saturated over time (Feely et al., 2004).

Figure 1.4. Distribution of (A) aragonite and (B) calcite saturation depth (Ω = 1) in the global oceans. The level at which aragonite and calcite are in thermodynamic equilibrium is known as the saturation depth. When the degree of saturation, Ω, is greater than 1, seawater is supersaturated with aragonite and calcite; conversely, seawater is under saturated with respect to these minerals when Ω < 1. This depth is significantly shallower for aragonite than for calcite, because aragonite is more soluble in seawater than calcite (Feely et al., 2004, 8p.2).

In the warm tropical and subtropical waters, under-saturation will occur when ρCO2 values reach ~1700 and ~2800 µatm, respectively (Kleypas et al. 2006). However in the cold high-latitude surface waters of the subarctic North Pacific and North Atlantic, and in the sub-Antarctic and Polar Regions of the Southern Ocean, aragonite and calcite under-saturation will occur when ρCO2 reaches ~600 and ~900 µatm, respectively (Kleypas et al., 2006). Figure 1.5 shows the decrease in saturation levels predicted globally for up to the year 2100, noting the extremely low saturation levels for polar surface waters, with high under-saturation predicted for 2100. It makes sense to be concerned how these chemical changes will affect organisms who live in the sea.

Figure 1.5 Estimated aragonite saturation states of the surface ocean for the years 1765, 1995, 2040, and 2100, this is an example for shallow and deep corals under predicted CO2 emission scenarios (Kleypas et al., 2006 p.17) .

1.3 Marine life in an acidifying ocean

Marine invertebrates and early life stages

Research into marine stressors, such as acidification, has been increasingly focused on understanding how marine species will respond to ocean change (Byrne, 2011). Under the predicted decreases in ocean pH, calcification will be the major biological process impacted (Dupont et al., 2010b), with calcifying marine organisms (e.g., molluscs, foraminifera, crustaceans, echinoderms, corals and coccolithophores) predicted to be most vulnerable to ocean acidification because calcification rates may decrease as a 2- result of reduced CO3 availability (Dupont et al., 2010b, Findlay et al., 2009a), the most important ion in this process. Ocean acidification is a particular threat to 2- calcifying pelagic organisms because it decreases availability of the CO3 required for the production of a skeleton (Byrne, 2011) (Figure 1.6). CaCO3 production is dependent on the ability to increase calcification thus counteracting dissolution (Findlay et al., 2009a). Inhabiting the pelagic euphotic zone are a diverse and often abundant community of coccolithophores, foraminifera and pteropods in most open and coastal ocean areas (Raven et al., 2005). Giant blooms of cocolithophores can produce one million tonnes of calcite (Holligan et al., 1993). Ocean acidification studies have 2− shown a decline of coccolithophore mass with decreasing CO3 (Beaufort et al., 2011). Similarly, this response is integrated with the predicted decrease in calcification of planktonic foraminifera (Moy et al., 2009). changes to the distribution and abundance of these protists could have significant effects on the global carbon cycle (Raven et al., 2005). Larger pelagic organisms are just as vulnerable; studies show that dissolution of the female brood chamber in the paper nautilus (Argonauta nodosa) could compromise their existence in a more acidified world (Wolfe et al., 2012).

Multiple studies have indicated that the calciﬁcation rates of tropical reef-building corals will be reduced by up to 60% when the CO2 concentration reaches 560ppm in 2050 (Gattuso et al., 1998, Kleypas et al., 2006, Raven et al., 2005). Many marine calcifiers also differ in their forms of the bio-mineral calcium carbonate (Lowenstam and Weiner, 1989), therefore responses to ocean acidification are expected to vary by taxon (Hofmann et al., 2008).

Calcium ions

2+ Ca

1µm Calcification

Figure 1.6 Absorbed CO2 in seawater (H2O) forms carbonic acid (H2CO3), lowering the water's pH level and making it more acidic. This raises the hydrogen ion

concentration in the water, and limits organisms' access to carbonate ions, which are needed to form hard parts. (modified from Kleypas et al., 2006, p.15; UKOA 2013)

Importance of larvae and juveniles

Developmental stages of echinoderms are the most sensitive and vulnerable part of the life cycle and any interruptions of this life stage will inhibit adult populations. Morphological abnormalities later in the life cycle could affect reproduction or even survival to a reproductive age. Larval performance ie. Growth and feeding ability, has been shown to remain constant at higher seawater CO2 concentration in adults and juveniles of active, high metabolic species with a powerful ion regulatory apparatus (Melzner et al. 2009). Thus, larvae and juveniles that lack advanced powerful ion regulatory systems may not be able to compensate for increased CO2 concentration. Yamada & Ikeda (1999) revealed that the tolerable pH range for marine plankton decreases with increasing exposure time, therefore non-significant early larval mortality rates may increase in the later stages of development; this decrease could be due to later failure of acid base regulation that was not noticeable at a morphological level. pH decrease could lead to acid-base up-regulation, which may come at a cost, impacting

11 the animals’ well-being, for example, increased muscle degradation in the brittle star Amphiura filiforms (Wood et al 2008). Depending on species and stressor type, early embryos and larvae exhibit different sensitivities to stressors. Successful recruitment and persistence of populations require that all ontogenetic stages are completed successfully (Gibson et al. 2011).

Many marine invertebrates broadcast-spawn their gametes for external fertilization and have pelagic larvae that spend from hours to months in the water column before settling in the adult habitat (Byrne, 2011, Dupont et al., 2010b) (Figure 1.7). Their high sensitivity to water chemistry has allowed them to be a good subject for monitoring environmental pollutants (Byrne, 2011). Earliest life history stages of shellfish larvae have been shown to be especially vulnerable to high CO2, displaying significant declines in survival and delays in metamorphosis at CO2 levels projected for later this century (Talmage and Gobler, 2010). In addition, juvenile benthic organisms may be more vulnerable to ocean stressors than the adults (Foster, 1971). Different life stages will have differing sensitivities to climate change stressors, yet compromised performance of any developmental stage results in a negative effect for adult populations (Harley et al., 2006). Table 1.2a shows examples of molluscs, pteropod, and foraminifera negatively affected by experimental lowering of pH. In contrast, the possibility for increased or maintained calcification under high CO2 conditions exists, 2- as there is evidence that some organisms do not rely solely on CO3 to calcify. Where most calcification levels are maintained or slowly decrease, some organisms have been shown to completely overcome dissolution and increase their calcification rate, for example increased growth of shell height and width with acidification of the marine snail Littorina littorea (Findlay et al., 2009a).

To date, studies of the effects of elevated CO2 have been largely focused on corals and algae (Doney et al., 2009, Hoegh-Guldberg et al., 2007). Yet there remains a lack of understanding in our knowledge of the physiological and ecological impacts of increasing ρCO2 on other benthic calcifiers such as echinoderms, molluscs and deep sea corals. A group with well-defined calcified life-history stages that is amenable to laboratory spawning and culture will be ideal for exploring these ideas further.

Figure 1.7. Schematic diagram of the life cycle of the seastar Patiriella regularis. Different life stages will have differing sensitivities to climate change stressors (modified from Byrne 2011, p.3).

Phylum Echinodermata

Echinoderms are among the most familiar seashore organisms widely distributed in all oceans at all depths. They are a vital component of the marine environment and found in almost every ecosystem, where they are often keystone ecosystem engineers (Dupont et al., 2010b). Their five subclasses are: crinoids (sea lilies and feather stars), asteroids (starfishes), ophiuroids (brittlestars), echinoids (sea urchins and sand dollars) and holothuroids (sea cucumbers). It is often hypothesized that echinoderms will be strongly impacted by ocean acidification and that differences will be observed between taxa (e.g. highly calcified sea urchins being more affected than less calcified sea stars) and stages (e.g. non-calcifying larval stages being less affected than calcifying adults in sea stars) (Dupont et al., 2010b). In order to successfully create a skeleton, echinoderms

13 require a continuous supply of Ca2+ ions to supersaturate the site of deposition. Without full calcification of vital skeletal structures, echinoderms may struggle to survive and be more vulnerable to lethal damage (Bentov et al., 2009, Pelletier, 2010).

Sea urchin calcification

The test plates of the sea urchin are made of one of the most soluble polymorphs of

CaCO3, high magnesium calcite (Mg-calcite), which is at least 50% more soluble in seawater than calcite on its own (Fabry et al., 2008, Mucci, 1983). High Mg-calcites with Mg greater than about 10 mol% (14 wt%) are known to be even more soluble than aragonite (Morse and Mackenzie, 1990). With these minerals being the most soluble common carbonate minerals, it is expected that marine organisms made from these calcites will be the “first responders” to a declining saturation state (Morse et al., 2007). Therefore the mineralogy of sea urchins makes them highly susceptible to dissolution at lowered pH (Beniash et al., 1997, Findlay et al., 2009a, Kobayash.S and Taki, 1969, Raz et al., 2003). Echinoderms larval skeletal elements (rods, adult test, teeth and spines) form via an amorphous calcium carbonate precursor stage, 30 times more soluble than calcite (Politi et al., 2004) thus adding to their vulnerability to future ocean acidification (Figure 1.8). Recent studies of the effects of increased ocean CO2 on sea urchins revealed smaller size and body weight under lowered pH, along with a thinning of the CaCO3 test (Clark et al., 2009, Gonzalez-Bernat et al., 2013a, Pecorino, 2012). Furthermore, shell damage, malformed skeletogenesis, decreased survival, reduced calcification and dissolution of larval skeleton have been observed (Table 1.2b). Larval size reduction is thought to impair performance and have negative consequent effects for benthic adult populations (Brennard et al 2010).

Figure 1.8 Evechinus chloroticus larvae under normal light microscope (left) and Arachnoides placenta larvae under polarised light, showing larval skeleton (right). Scale bars = 100µm. 14

Table 1.2 Images of the physical effects of increased seawater CO2 on a variety of marine organisms, including species of a) mollusc, pteropod, foraminifera and b) echinoderms.

Ambient Conditions Increased seawater CO2 Source a. Molluscs, pteropod, foraminifera (Talmage and Gobler, 2010) Hinge from 36 day old Mercenara mercenara larvae

(Bednarsek et al., 2012) Limacina helicina antarctica larvae

(Riebesell et al., 2000) Gephyrocapsa oceanica scale bar =1µm

(Kurihara, 2008) Mollusc Crassostera gigas scale bar = 50 µm

(Byrne et al., 2011) Haliotis coccoradiata larvae (at 20°C)

b. Echinoderms (Dupont et al., 2008) (Ophiothrix fragilis larvae scale bar = 10µm)

(Clark et al., 2009) (Evechinus chloroticus larvae scale bar = 5µm)

Ambient Conditions Increased seawater CO2 Source 15

Ambient Conditions Increased seawater CO2 Source

(Clark et al., 2009) (Pseudechinus huttoni , scale bar = 5µm)

(Kurihara et al., 2004) (Echinometra mathaei, scale bar = 50µm )

(Kurihara, 2008)

(Hemicentrotus pulcherrimus , scale bar = 50µm)

(Byrne et al., 2011) Heliocidaris erythrogramma newly metamorphosed 5-day- old juveniles.

(Brennand et al., 2010) 5 day old Tripneustes gratilla larvae

(Gonzalez-Bernat et al., 2013b) 4 day old Arachnoides placenta

(Gonzalez-Bernat et al., 2013a) 58 day old Odontaster validus

Tropical, temperate, polar echinoderms

Latitudinal trends in growth, size, and skeletal composition of invertebrates are extremely useful in the context of understanding ocean acidification (Smith and Lawton, 2010). Low-latitude echinoderms have shown signs of stress to a decreased ocean pH, for example, subtropical juvenile echinoderms stopped growing and produced more brittle and fragile exoskeletons in acidified seawater (Shirayama and Thornton, 2005). Although tropical species have shown negative effects to decreased ocean pH, it is understood that polar to temperate waters are likely to experience the full impact of ocean acidification due to low water temperatures and high rates of physical overturning increasing uptake of CO2 (Raven et al., 2005). Cold high-latitude waters are the first to become carbonate under-saturated (Byrne, 2011), so ocean acidification is potentially a serious stressor for polar species. High-latitude regions 2- such as Antarctica and the Arctic have naturally lower CO3 concentrations compared to lower latitude regions due to increased CO2 solubility, thus echinoderms which live in these cooler waters exist under conditions less favourable for calcification than those living at lower latitudes. Additionally the inclusion of magnesium calcite to their skeletons adds to the corrosion stress (Andersson et al., 2008). Polar seawater saturation states with respect to CaCO3 minerals is thus lower than in tropical or even temperate regions (Fabry, et al. 2009). Recent ocean acidification studies have shown poorly-developed Antarctic pteropods, foraminifera and sea urchins (Bednarsek et al., 2012, Comeau et al., 2009, Gonzalez-Bernat et al., 2013a, Moy et al., 2009). Pteropods form shells made of aragonite, a metastable polymorph of CaCO3, which like high Mg- calcite is prone to dissolution (Fabry et al., 2008). Secondly, when living Clio pyramidata were exposed to a level of aragonite undersaturation (projected for Southern Ocean surface waters in year 2100 under the IS92a emissions scenario) there was marked shell dissolution within 48 h (Feely et al., 2004, Orr et al., 2005).

Larvae and juveniles that lack advanced powerful ion regulatory systems or have a slow metabolism, such as Antarctic species Sterechinus neumayeri, may not be able to cope with change, which only compounds the effects of an environment which already has elevated levels of CO2 (Clark et al., 2009). Polar animals act as a natural laboratory for understanding the impacts of climate change and in collaboration with the low CO2 saturation, act as a bellwether for other marine systems (Aronson et al., 2007).

Seasonal conditions also need to be taken into consideration as wintertime ocean warming has been observed in south-east Australia, thus potentially affecting winter spawners to a greater degree, as well as planktonic phases (Poloczanska et al., 2007). In the Southern Ocean there is a decrease in CaCO3 saturation, with the winter of 2030 expected to be undersaturated with respect to aragonite (Figure 1.9) (McNeil and Matear, 2008). Impacts of ocean change on marine invertebrate reproduction and development need to be considered in a regional context. A taxonomically broad effort, including a variety of calcifiers, will capture individual responses that can be integrated to reveal impacts on ecosystem-level processes.

Figure 1. 9 Seasonal estimates of pH and CO32- for the Southern Ocean. (Top) Winter and summer distributions of pH. (Middle) Winter and summer distributions of CO32- (McNeil and Matear, 2008, p.2).

1.4 Acid-base regulation

Many experimental studies have demonstrated that elevated seawater CO2 can have a sub-lethal impact on an organism’s development and physiological features, especially in marine calcifying organisms (Doney et al., 2009, Fabry et al., 2008, Guinotte and Fabry, 2008). Much less is known about the cellular-level mechanisms (e.g., acid base regulation), that alter these processes in response to ocean acidification.

Why is acid-base regulation important?

The acid–base status of an animal is affected by a range of functional and environmental stresses (e.g., ocean acidification, ocean warming); the change depends upon the effects of these stressors on the mode and rate of metabolism, respiration, and the mechanisms of H+ equivalent ion exchange (Reipschläger and Pörtner, 1996). Extracellular acid–base regulation plays an important role in calcification rates of many marine calcifiers; it also influences other physiological processes (such as ion pumping) as many are strongly pH dependant (Brownlee, 2009). Intra- and extracellular pH can alter a cell's functioning, because it is important for the activity of a number of enzymes which are pH sensitive, as well as for the efficiency of contractile elements and the conductivity of ion channels (Johnson et al., 1976, Madshus, 1988). Acid–base status is not only operative in calcifiers but also non-calcifiers in setting CO2 sensitivity (Portner et al., 1998). Biomineralization can be intracellular (e.g., coccolithophoriids) or extracellular (most of the foraminifera and many other invertebrates) (Bentov et al., 2009), thus acid–base parameters, especially pH, can affect this process as well as other metabolic processes. Studies by de Nooijer et al. (2008) found that foraminifera elevate their pH at the site of calcification to increase precipitation of their very low Mg- calcite, which helps to increase the incorporation of Mg2+ into calcite. In contrast, some organisms are able to maintain pH homeostasis with cytosolic values lower than the surrounding seawater, such as the Cnidarian Anemonia viridis (Brownlee, 2009).

Extracellular and intracellular pH regulation in developing echinoderms

Understanding acid–base regulatory capacities in echinoderms is important because formation and maintenance of the calcium carbonate skeleton is dependent on pH homeostasis (Stumpp et al., 2012a), which is carried out by transport proteins situated in the plasma membrane (Baltz, 1993). Many processes of echinoderm development, especially larval development are initiated by an increase in intracellular pH (pHi), such as the activation of egg by sperm, where pHi is raised permanently by 0.3 units resulting from an exchange of extracellular Na+ for intracellular H+, via the sodium coupled exchange (Johnson et al., 1976). Furthermore, sperm pHi increases upon dilution of semen into seawater. For a sufficient increase to trigger sperm pHi must be greater than 7.2 (Johnson et al., 1983). The induction of the acrosome reaction also requires an increase in pHi to ~7.6 (Schackmann et al., 1981). pHi affects protein synthesis, protein phosphorylation, polymerization of cortical actin (Payan et al., 1983), all of which are important in the development of echinoderms. pH oscillations are important in controlling the cell cycle and the proliferative capacity of cells (Madshus, 1988). Additionally, all cells have a Na+/H+ antiporter, important for the regulation of pHi after acid loads. Acid–base parameters, especially pH, may affect cell functions and metabolic processes (Reipschläger and Pörtner, 1996). For example, metabolic activation results from the change in pHi during egg activation which can be prevented by lowering the extracellular pH (pHe) below 6.5, preventing acid efflux (Johnson et al., 1976). The production of a skeleton relies on the internal regulatory capacity of echinoderms, calcification occurs in isolated compartments where ion transport across various epithelia establishes an environment suitable for calcification to occur (Portner, 2008).

Echinoderms are heavily calcified and can have two quite different and contrasting episodes of skeletogenesis, one during the larval phase and the other during the adult phase (Dupont et al., 2010b). In adult echinoderms, skeletal ossicles are produced within an extracellular syncitium formed by associations of scleroblastic cells. In larvae, skeletogenesis takes place within a compartment whose boundary comprises a syncitium of primary mesenchyme cells (PMCs) (Dupont et al., 2010b). These PMCs are located within the extracellular matrix of the primary body cavity, where they form a syncytium around the growing spicules (skeleton rod) in pluteus larvae (Figure 1.10).

PMC’s produce skeleton in a complex species-specific manner (Madshus, 1988), in which the enzyme carbonate anhydrase, an enzyme that drives CO2 elimination in cells, catalyses the reaction in which CO2 and H2O are converted to H2CO3 (Mitsunaga et al., 1986). This syncytial sheath covers the entire surface of the spicules and communicates with the extracellular environment of the primary body cavity (Stumpp et al., 2012a). This complex mechanism is very important for calcium deposition in the spicules. If a suitable environment cannot be established and maintained, development of the larval skeleton will be prohibited, perhaps to a degree that inhibits development into an adult (Politi et al., 2004, Stumpp et al., 2012a).

Figure 1.10 A cross section of a sea urchin spicual during biomineralization. 1, various trans-membrane transporters for HCO3- and calcium from surrounding; 4, action of carbonic anhydrase (CA), an enzyme that converts CO2 in the cell; 5,certain spicule matrix proteins (SM) are involved in directing biomineralization in the extracellular space between the PMC and the growing skeleton (Hofmann et al., 2008, p.4).

How will ocean acidification affect pHe and pHi?

The capacity to regulate and compensate for acid-base disturbances is crucially important in setting sensitivity to hypercapnia (Portner et al., 1998). Extracellular fluid (in conjunction with intracellular fluid) helps control the movement of water and electrolytes throughout the body. It is the first compartment affected by water

21 physiochemistry and responds in a species-specific manner acting as a mediator of the effects that water chemistry has on a species (Portner, 2008). Decreases in larval development and delays in metamorphosis may result in a higher mortality during the planktonic life phase (Dupont et al., 2010b). Predicted ocean decreases in pH could have multiple effects on the ability of echinoderms to regulate internal parameters such as pHe and pHi. To understand the full effects of ocean acidification on echinoderms and determine factors that interrupt larval development, settlement, calcification and other physiological processes need to be investigated in the organism with respect to internal parameters such as pHe and pHi.

Decreased seawater pH could make it more difficult to maintain a suitable environment not only for calcification but for enzyme reactions, needed in everyday cell homeostasis. Species that lack a good internal regulatory system may be more vulnerable to ocean acidification. For example the brittle star, Amphiura filiformis, has a poor internal regulatory capacity (Findlay et al., 2009a). When exposed to lower seawater pH the internal fluids have a similar chemical composition to the surrounding seawater, bathing the endoskeleton and inner shell surface in low pH seawater causing measurable decreases in calcification.

It is a very energetically costly undertaking for PMC acid–base regulation because PMC’s have to maintain full cell functionality in a more acidified extracellular medium (Stumpp et al., 2012a). It has also been noted organisms without extracellular fluid

(unicellular organisms and gametes) are more vulnerable to elevated levels of ρCO2, while the presence of extracellular fluid may dampen the effects of seawater pH decrease in the intracellular fluid by compensation (Melzner et al., 2009). Another study has demonstrated that metabolic depression during steady-state hypercapnia is mediated by a decrease in extracellular pH (Reipschläger and Pörtner, 1996). Yet, some species have the ability to compensate for pH changes by the accumulation of bicarbonate in mostly extracellular (but also intracellular) compartments.

Extracellular bicarbonate accumulation will support compensation of intracellular acidosis through trans-membrane ion exchange (Portner et al., 1998). For example, (Lindinger et al., 1984) reported that hypercapnia caused respiratory acidosis in the extracellular fluid while the intracellular space had a higher buffering capacity in the

22 bivalve Mytilus edulis. Similar buffering also occurred via accumulation of bicarbonate in the extracellular space sufficient to compensate for hypercapnia-induced intracellular acidification in the marine worm Sipunculus nudus (Portner et al., 1998). This buffering process provides a preconditioning for the re-establishment of the control of intracellular pH. Sea urchins (except cidaroids) have a higher buffer capacity in their inner extracellular fluids than that of seawater and increase this capacity at low pH. Their respiratory metabolism is either increased or unaffected. On the contrary, starfish have a buffer capacity in their inner fluids similar to that of seawater and decrease their metabolism at low pH (Collard et al., 2013).

Sea urchin larvae raised under acidified conditions showed increases in the enzyme NKA (α-subunit), which provides an electrochemical gradient for most secondary active transport processes (Martin et al., 2011, Stumpp et al., 2011b). This up- regulation of NKA results in an increase in metabolic rate, indicating a higher energy demand (Stumpp et al., 2011a). Additional acid base stress, such as calcification during environmental hypercapnia, significantly impacts an organism’s energy budget (Stumpp et al., 2012a). This change in energy could reveal that energy is not focused purely on growth and calcification, but more on trying to regulate and maintain pH, which may be why acidified echinoderm larvae were stunted in previous ocean acidification experiments (Table 1.2). Stumpp et al. (2012) suggest most energy is allocated to PCM acid base regulation because these cells need to maintain full function in acidified waters. Therefore, it is important to measure pHe changes in different larval regions and understand how it is affected by changes in seawater pH. For example, measuring pHe surrounding the primary mesenchyme cells allows evaluation of the potential impact on skeletal development. Figure 1.11 illustrates the process of calcification and pH regulation. A decrease in pHe directly affects the calcifying primary mesenchyme cell (PMC) syncytia challenging the intracellular pH (pHi) regulatory machinery due to decreased proton gradients.

Figure 1.11 Schematic model summarizing the interplay of calcification and pH regulation. pH homeostasis is maintained by ion transporters, which either directly or indirectly depend on the consumption of energy. Increased energetic costs due to decreased proton gradients lead to a shift in the larvae’s energy budget, which decr eases scope for growth and may also translate into juvenile impaired fitness. ST, stomach. (Stumpp et al., 2004, p.4).

Gene expression in the PMCs is thought to be involved in calcium transport; calcium is transported from the external seawater, modified in the PMC cytoplasm, and then moved via exocytosis into the extracellular space around the forming spicule (Hofmann et al., 2008). Dupont et al. (2010b) believes this mechanism of calcite deposition (matrix secretion, ion transport pathways) will follow similar cellular and molecular pathways in both larval and adult life stages, therefore, impacts of environmental perturbations, such as pH, might be equally important in both adults and larvae. pHe is important for cell function with most enzymes being pH specific. Larval echinoderm stages are the most sensitive and vulnerable phase, additionally, due to latitudinal differences polar species live in a naturally higher CO2 environment and extremely cold waters resulting in very slow metabolism rates. The polar sea urchin, Sterechinus neumayeri takes at least 20 days to reach the pluteus (4 arm) stage (Clark et al., 2009), whereas the tropical sand dollar, Arachnoides placenta only takes 24 hours (Poloczanska et al., 2007). It is clear that pH tolerance deserves more investigation in echinoderms, especially how it influences extracellular systems and lifecycles of marine organisms considered in a multi-latitude context.

1.5 Aims and objectives

To understand the potential effects of ocean acidification we need to investigate variation in parameters such as extracellular pH (pHe) change in developmental stages of marine species when exposed to reduced seawater pH.

The main objectives of this thesis are to:

1. Develop and apply a technique to measure pHe in echinoderm larvae that is non-toxic, non-invasive and can be experimentally tested across various pH treatments and species.

2. Determine the effect that predicted future ρCO2 seawater concentrations will have on the pHe of echinoderm developmental stages, in particular, examine changes in : a. pHe at different stages of development (gastrula, pluteus and seastar bipinnaria development) b. pHe differences across different regions of the 4 arm pluteus (arms, gut, oesophagus, oral hood)

3. Determine if there is a broad latitudinal difference (tropical, temperate and polar) in sensitivity to decreased seawater pH by looking at five species ranging from: Arachnoides placenta (tropical), Evechinus chloroticus, Pseudechinus huttoni (temperate), Sterechinus neumayeri, and Odontaster validus (polar).

Hypothesis 1: As CO2 ppm increases, echinoderm larvae will be less able to regulate their pHe as seawater pH decreases. Acid base regulation will be up-regulated to actively maintain a favourable pHe, but this change in energy allocation will cause failures in other vital areas. Ultimately larvae will show a decreased ability to regulate pHe under higher ρCO2 conditions.

Hypothesis 2: Echinoderm species from polar regions will be more sensitive to pH change than those from warmer waters. Due to their slow metabolism and long larval life cycle they will be less capable of regulating extracellular pH.

The following chapters include:

Chapter 2. Measuring Extracellular pH (pHe)

A detailed chapter of how the method to measure extracellular pH was developed using the probe 8-hydroxypyrene-1,3,6-trisulfonic acid (HPTS) in echinoderm larvae raised under various CO2 ppm conditions.

Chapter 3. Response of Extracellular pH (pHe) to Reduced Seawater pH in Embryos and Larvae of Five Echinoderm Species

Report of the experimental results of decreased seawater pH for the experimental predicted future pHs of pH 7.8, pH 7.6 and pH 7.4 by evaluating morphometrics and extracellular pH on the early development of the temperate sea urchins Evechinus chloroticus and Pseudechinus huttoni from Fiordland, New Zealand. The tropical sand dollar Arachnoides placenta, collected from Townsville, Australia. Also, the experimental results of decreased seawater pH for the treatments of pH 7.7, pH 7.5 and pH 7.2 by evaluating the morphometrics and extracellular pH on the early development of two ecologically important Antarctica species, the sea urchin, Sterechinus neumayeri and asteroid, Odontaster validus.

Chapter 4. General discussion and Conclusion

Discussion of overall findings and conclusions for this study.

Chapter 2 Measuring extracellular pH (pHe)

2.1 Introduction

Methods of measuring Extracellular pH

There are several methods which enable the measurement of intra- and extracellular pH (pHi and pHe respectively) in a cell. Most commonly used for pHi measurements are H+-selective microelectrodes or the incorporation of highly charged pH sensitive fluorescent compounds into cells. The methods to import these compounds into cells include microinjection (Oparka et al., 1991), scrape loading (Giuliano and Gillies, 1987), and hypertonic lysis (Green, 1988). Although all of these approaches have the disadvantage of perturbing the cell resting state physiology (Han and Burgess, 2009). Fluorescent dyes have the potential to help visualize pH within organisms without disturbance. Previous studies by Stumpp et al. (2012) used a non-invasive method of the most widely used pH fluorescent dye, 2’7’-bis-(2-carboxyethyl)-5-(6- carboxyfluorescin) or BCECF, to investigate pH within the extracellular matrix (ECM) in sea urchin larvae. BCECF uses a dual excitation (λ1 = 490 nm and λ2 = 440 nm) and fixed emission at 535 nm. Using ratiometric fluorometry between the two wavelengths pHi can be estimated within the cell (Stumpp et al., 2012a). Here, BCECF was taken up into the gut and exported from the cytosol of cells lining the gastric tract and subsequently trapped in the ECM of the body cavity. BCECF's optimum pH range in seawater is between pH 6.5 and 7.5. This inability to measure fluorescence below pH ~6 limits quantitative pH estimation in more acidic compartments (pH <6) (Han and Burgess, 2009, Probes, 2006), limiting the overall utility of the dye. Intracellular hydrolysis of the ester bond has also been shown to cause significant intracellular

27 acidification over time (Aharonovitz et al., 1996). Additionally, the excitation or emission spectra of BCECF does not shift with pH, limiting the dynamic range of fluorescence ratio determinations (Gan et al., 1998). The probe is also very susceptible to photo-bleaching (Han and Burgess, 2009) leading to progressive deterioration of the signal and damage of the biological preparation, likely associated with the generation of oxygen radicals (Gan et al., 1998). BCECF also has the disadvantage that its fluorescence emission intensity is dependent on the concentration of the probe, so if the dye accumulates regionally, it can indicate false internal pH values which are due to dye concentration, not proton concentration differences (Han and Burgess, 2009). Another ratio-imaging fluorescent probe which has been used to measure pHi during sea urchin egg activation is a seminaphthorhodafluors dye, carboxy SNARF-1. Visible light–excitable fluorescent pH indicators (Suzuki et al., 1995) are useful for measuring pH changes between pH 7 and pH 8 (Probes, 2003). What is needed for ocean acidification research, however, is a dye that can indicate a wider range of pH, such as 8-hydroxypyrene-1,3,6-trisulfonic acid (HPTS).

HPTS and its applications

HPTS is a non-toxic highly water-soluble fluorescent pH indicator. Excitation ratio exc exc imaging is possible, with absorbance maxima at 405 nm (λ1 ), and 460 nm (λ2 ). The absorption at each wavelength is pH dependent as the solution is varied from pH 5 to 8 (Figure 2.1). It has a single emission optimum at 520 nm (de Nooijer et al., 2008, Han and Burgess, 2009, Wolfbeis et al., 1983). HPTS has been used for measuring intracellular pH distributions in plants (Wright and Oparka, 1996), mammalian cells (Overly et al., 1995), and more recently, to study the pH distribution in several species of intact foraminifera (de Nooijer et al., 2008), even though HPTS is a membrane impermeable dye that does not usually label the cytosol via endocytosis. HPTS has been used for measurement of cytoplasmic pH and acidic organelle pH in many cell types (Han and Burgess, 2009), by means of microinjection, electroporation (Pena et al., 1995) or liposome-mediated delivery (Pena et al., 1995, Spragg et al., 1997).

HPTS has many properties that make it a useful probe for measuring pHe: 1) it is non- toxic and does not interfere with the natural pre-functioning pH at experimental conditions (Willoughby et al., 1998); 2) it has a near-neutral pKa and high pH

28 resolution in the physiological range; 3) it responds rapidly to pH changes; 4) it is a ratiometric indicator, allowing quantitative measurements regardless of organelle size or probe concentration; 5) it can be calibrated within live cells, for precise and accurate quantification; 6) it is hydrophilic and membrane-impermeant, allowing easy loading into endosomes by fluid-phase endocytosis but preventing leakage across intracellular membranes (Overly et al. 1995) and 7) as noted by Wolfbeis et al. (1983) HPTS fluorescence was stable from pH 0 to pH 14. HPTS is thus an ideal fluorescent indicator for the measurement of pHe. This non-invasive form of measuring pH has had little use in examining how ocean acidification affects acid-base regulation and pHe homeostasis in marine organisms. However, by incubating echinoderm larvae in this pH probe, we can visualize pHe in developing larvae, investigating acid base regulation when larvae are reared under varying seawater ρCO2 conditions.

Aims The objective of this chapter is to develop a technique and software to apply to living echinoderm larvae which will provide measurements of extracellular pH.

2.2 Methods 2.2.1 Fluorometry Absorption/Emission and pH of HPTS

Changes in fluorescence colour or intensity with pH is usually due to reversible dissociation processes, not only effecting the fluorescence spectrum but the absorption spectrum aswell (Figure 2.2) (Wolfbeis et al., 1983). HPTS exhibits a pH-dependent absorption shift from 405 nm (λ1 exc) to 460 nm (λ2 exc) as pH increases (Figure 2.1). It shows a maxima absorption at 460 nm in alkaline solution, which declines when the solution becomes acidic (Wolfbeis et al., 1983). When excited at 405 nm the reverse can be seen with greater absorbance in the more acidic solution (Figure 2.2). Only the phenolate (base form) is able to absorb light, when excited at 450 to 490 nm. Therefore, the more acidic the solution becomes, the less light will be absorbed and consequently also emitted (Wolfbeis et al., 1983).

1M NaOH

1M HCL

Figure 2.1 pH dependence of the absorption spectrum of HPTS . Maximum absorbance is shaded red ~405 nm (λ1 exc) , ~460 nm (λ2 exc ). λ (nm) on the x-axis is the excitation wavelength and ‘E’ (y-axis) = emission (modified from Wolfbeis et al., 1983).

2.2.2 Measuring the difference in emission/absorption Calibration pH is measured by calculating the ratio of the emission from the two wavelengths (λ2 exc exc /λ1 ). To relate this ratio to the larval developmental stage in seawater, a 12 point

30 calibration curve was carried out before examination of pHe in each species. 1 L of 30 µM HPTS in seawater solution was made and the pH was adjusted by the addition 1 M of HCl or NaOH and allowed to mix on a magnetic stirrer until a stable pH was reached exc exc (Figure 2.2). Non-seawater calibrations of HPTS (λ1 /λ2 ) have been shown to be independent of HPTS concentration within a range of 20 to 40 µM (de Nooijer et al., 2008, Overly et al., 1995). Photographs were taken of a small drop of solution on a slide from 12 pHs ranging between pH 4 and pH 10, with two samples photographed for each calibration pH. Samples were imaged using a Nikon fluorescent microscope Eclipse TE300 fitted with a Nikon filter cube (exciters UV1A and B2A, band pass filter HQ535/50) and plan fluor dry objectives (20x, 40x and 100x). Exposure time was set at 2 seconds for each image.

1M HCL

1M NaOH

Figure 2.2 Plot of HPTS fluorescence intensity at emmision 520 nm as a function of the pH-value at various excitation wavelengths. When pH is alkaline the HPTS solution is bright green/yellow and fluorescence intensity is high when excited at 454 nm (right), this disappears gradually as the pH is lowered and becomes more alkaline (left), fluorescence intensity also shifts and becomes greater at 403 nm (modified from Wolfbeis et al., 1983). 31

Fluorescent image processing was carried out using the computer programme MATLAB. The emission (520 nm) was photographed after excitation at the dual wavelengths 405 nm and 460 nm. Using the red, green, blue (RGB) scale, the ratio of 'green' light intensity (on a scale of 0 to 255) in the two images was calculated via a exc exc matrix for each pixel in the image (λ2 /λ1 ) (Figure 2.3). For each species, a 12 point ratio vs. pH calibration was graphed on Microsoft Excel to create a polynomial curve (Figure 2.4). A non-linear regression was calculated for each curve (if R2 was >0.95), and imported into MATLAB code for the measurement of pHe values in larvae (Figure 2.5). For each species a new standard curve was produced to eliminate the effects of different water temperatures, water chemistry and the use of different fluorescent microscopes.

Excitation

460nm 405nm pH Ratio (λ2 exc) (λ1 exc) λ2 exc/λ1 exc

9.44 0.94

6.18 0.4

3.34 0.2

Figure 2.3 An example of fluorescent images and ratios produced by three pH calibrations from the addition of 1 M of HCL and NaOH to 30µM HPTS seawater solution. Excited at wavelengths 460 nm (λ2exc) and 405 nm (λ1exc), and photographed at 520nm using a set exposure of two seconds.

9 P. huttoni y = 12.52x3 - 18.544x2 + 9.1582x + 5.1608 8.5 R² = 0.9776

7.5

6.5 pH (NBS)

5.5

4.5 0 0.2 0.4 0.6 0.8 1 Ratio λ2/λ1

Figure 2.4 Example of a calibration curve for 30µM HPTS. Equation and R2 value of the polynomial. Note only eight points shown as readings above pH 8.5 become saturated at a ratio of 1 and below 5 a ratio of 0 (no florescence) so readings at these values were removed.

2.2.3 Larval photo analysis

Incubation of Larvae and Extracellular pH measurements

Fertilised echinoderm eggs were developed under a range of seawater CO2 conditions (see chapter 3). During different stages of development, larvae were incubated in a concentration of 30µM HPTS seawater in a controlled temperature room and time period dependent on species (described in detail later in chapter 3). Embryos and larvae were rinsed several times in HPTS free seawater (in a Bogorov counting chamber) before photographing.

pH false colour mapping

The fluorescence of HPTS from larvae was photographed at a single emission (520nm) exc exc following excitation at both 405nm (λ1 ) and 460nm (λ2 ) consecutively. Using the exc exc standard curve, λ2 and λ1 images were divided pixel by pixel (as described above) resulting in a composite image of a false colour map of pHe (Figure 2.5 for code). Embryos were photographed using a set exposure of 2 seconds. Figure 2.6 shows images after both excitations and the false pH colour map produced in MATLAB.

Specific regions in the larvae (gut, larval rods, and oesophagus) were isolated in MATLAB (step 4, Figure 2.5) before calculating the average pHe in an area that was typically 50µm2 (Figure 2.6).

clear all; pic1=imread('blue1.tif','tif'); figure; image(pic1) impixelinfo axis equal pic2=imread('green1.tif','tif'); figure; image(pic2) impixelinfo 1 axis equal p1=pic1(:,:,2); p1=double(p1); p2=pic2(:,:,2); p2=double(p2);

%sel1=(p1(:,:)<1); sel2=(p2(:,:)<1); %p1(sel1)=NaN; p1(sel2)=NaN;

g=double(p2)./double(p1); 2 %f=(10.92*(g.^3)) - (16.78*(g.^2))+(8.709*g) + 5.176;%E.chloroticus %f=(3.1447*(g.^3)) - (10.119*(g.^2)) + (11.449*g) + 2.2453;%A.placenta %f=( 24.711*(g.^3)) - (39.175*(g.^2)) + (20.113*g) + 2.9342;%O.validus %f=(12.046*(g.^3)) - (19.349*(g.^2)) + (11.159*g) + 4.2428;%S.neumayeri %f=(12.25*(g.^3)) - (18.544*(g.^2)) + (9.1582*g) + 5.1608;%P.huttoni

clims = [ 6 9 ]; imagesc(f,clims); colormap jet colormap (flipud (colormap)); 3 colorbar; ylabel(colorbar,'pH');

impixelinfo

[x y]=meshgrid(1:size(f,2),1:size(f,1)); figure imagesc(x(:),y(:),f,clims)

[x2 y2]=ginput

[IN ON]=inpolygon(x,y,x2,y2); 4

ph=nanmean(f(IN(:,:)));

med=nanmedian(f(IN(:,:)));

Figure 2.5 MATLAB code used to produce pHe profile of larvae. 1) MATLAB reading images 405nm (blue) and 460nm (green) 2) Calibration equations for each species, from standard curves 3) pHe profile with limits between pH 6-9. 4) Allows manual isolation of regions and calculates the average pH within that region. See

MATLAB manual for explanation of coding.

Visible light 405nm

460nm pH colour map

O GW G

Figure 2.6 An example of a 1 day old A. placenta larvae imaged under (from top left); visible light, excitement at 405 nm, 460 nm and imaged at 520 nm, and the MATLAB generated pH colour map. The red dashed lines indicate where average pH measurements were made during the pluteus stage, this included the gut (G), larval rods (R), oesophagus (O), and in some cases the gut wall (GW) if it was defined.

Chapter 3 Response of Extracellular pH (pHe) to Reduced Seawater pH in Larvae of Five Echinoderm Species

3.1 Introduction

3.1.1 General latitudinal trends in seawater carbonate chemistry

CaCO3 saturation states of seawater have been shown to be highest in shallow, warm tropical waters with the levels lowest in cold high-latitude regions and at depth. Surface seawater saturation state with respect to carbonate minerals decreases with increasing 2– latitude owing to decreasing CO3 concentration and temperature (Feely et al., 2009). 2- Figure 3.1 shows variation in [CO3 ] with latitude and predictions of seawater pH and 2- [CO3 ], projecting a pH of 7.8 and aragonite undersaturation in Antarctic waters for 2- the year 2100. Low CO3 in the Southern Ocean is due to low surface temperatures and

CO2-system thermodynamics, additionally the upwelling of a large volume of deep water of the Antarctic divergence, containing a high [CO2 (aq)] concentration from organic matter remineralisation (Orr et al., 2005). The effect of these two together 2- result in a high positive correlation of present day [CO3 ] with temperature (Orr et al., 2- 2005). Relative to pre-industrial times, surface [CO3 ] has reduced by more than 10%, resulting in a reduction of 29 mmol/kg in the tropics and 18 mmol/kg in the Southern Ocean (Orr et al., 2005). Tropical and subtropical surface seawater is currently supersaturated with respect to calcite and aragonite by 550% and 360%, respectively.

Temperate surface waters are supersaturated with respect to these mineral phases by 370% and 230%, respectively, and high latitude polar surface seawater is supersaturated by 260% and 160%, respectively (Andersson et al., 2008).

2- Figure 3.1 Increasing atmospheric CO2, decreasing surface ocean pH and [CO3 ] over time and latitude predicted for the year 2100, (a) lead to reductions in surface ocean pH 2- (b) and surface ocean [CO3 ]. Results are given as global averages from 1994 data and the preindustrial (‘Preind.’), and include two future CO2 scenarios (IS92a (dotted line) ‘continually increasing’ scenario of 788 ppm in the year 2100 and S650 (thick dashed line) ‘stabilization’ scenario of 563 p.p.m.v. in the year 2100). The thin dashed lines 2- indicate the [CO3 ] for sea water in equilibrium with aragonite and calcite are nearly flat, revealing weak temperature sensitivity (Modified from Orr et al 2005 p.682).

The warm surface waters of tropical and subtropical waters may not become undersaturated with respect to calcite or aragonite by 2300 according to Feely et al.

(2009). However, some models indicate that even if atmospheric CO2 is stabilized at 450 ppm, only 8% of coral reefs will be surrounded by seawater that supports calcification (Cao and Caldeira, 2008), thus, tropical calcifiers could be particularly affected, furthermore they may be heavily impacted by the synergistic effects of acidification coupled with ocean warming, which will also be experienced globally (Brennand et al., 2010, Byrne, 2011, Byrne et al., 2011, Byrne et al., 2010b). In some regions associated with upwelling, shoaling aragonite saturation horizons may reach the depth ranges of calcifying benthic and pelagic species (Feely et al., 2004).

Temperate waters are expected to encounter undersaturation of aragonite by the year 2300 (Feely et al., 2004). Additionally, temperate waters experience high seasonal water temperature variation, consequently they could experience seasonal undersaturation of CaCO3, which is projected to occur in some areas of the Southern Ocean (McNeil and Matear, 2008). In this study our temperate species are from New Zealand waters, which lie at 45° south, is in close proximity to the Southern Ocean (50° S). They experience cold subsurface, sub-Antarctic currents (Raven et al., 2005, Carter,

1998), vulnerable to the low CaCO3 saturation conditions that the Southern Ocean experiences (Fabry et al., 2008). Recently, CO2 measurements from a 15-year transect in New Zealand's Exclusive Economic Zone show that surface ocean pH and carbonate availability are decreasing, as observed in sub-Antarctic waters (Currie et al., 2013).

High-latitude regions are expected to bear the brunt of ocean acidification with surface waters of polar and sub-polar regions projected to become undersaturated with respect to aragonite by the year 2040 - 2050 (McNeil and Matear, 2008). pH in Antarctic waters are relatively stable when compared to other sites in a recent study (Hofmann et al., 2011), yet there are still limited data for the Southern Ocean. The size of areas with aragonite saturation levels of 3.5 or higher are predicted to decrease to less than 5% by the end of the 21st century. By that time, large parts of the Southern Ocean will experience conditions that may lead to the dissolution of skeletons and shells due to the undersaturation of aragonite, projected to occur by 2100 (Orr et al,. 2005). If CO2 emissions are not stabilised the entire Southern Ocean may become undersaturated; open-water areas in this region already experience a strong seasonal shift in seawater

39 pH (0.3–0.5 units) between austral summer and winter, and wintertime aragonite undersaturation is projected to occur as early as 2030 (McNeil and Matear, 2008, McNeil et al., 2011).

Calcification variation with latitude

There is a well-defined gradient in the calcification of shells with latitude due to the availability of calcium carbonate in the ocean (Graus, 1974). Gastropods respond directly to the availability of calcium carbonate in the ocean with average calcification decreasing from low to high latitudes, leading to thinner shells in polar regions (Graus, 1974). Cold-water pelecypods commonly have small, thin and chalky shells; they are also missing spines and ornamentation (Nicol, 1967). At high latitudes, shell-building materials (Ca2+) are more difficult to remove from seawater because of the saturation state of CaCO3 (Zeebe, 2012). Increased dissolution has also been noted at cooler temperatures with the outer surfaces of bivalve shells exposed to pH 7.4 seawater showing deterioration after 35 days, and by 56 days it was possible to detect the exposure of aragonite or calcite prisms in the outer shell surfaces of the brachiopod Liothyrella uva and the bivalve Yoldia eightsi (McClintock et al., 2009).

Physiological responses to latitude

It is widely accepted that growth and metabolism are decreased in organisms living at higher latitudes (Fabry et al., 2008, Peck, 2002). Polar marine invertebrates often exhibit low metabolic rates (Peck, 2002, Peck and Conway, 2000), and very slow development and growth when compared to their temperate and tropical equivalents (Arntz et al., 1997, Pearse et al., 1991). Contrasting studies have found oxygen consumption, mitochondrial densities and mitochondrial capacities to be higher in polar ectotherms, concluding that invertebrate life is more costly at these higher latitudes (Peck, 2002). Physiological studies have revealed that echinoderms and bivalve molluscs may be some of the most vulnerable groups to ocean acidification because they are poor iono-regulators and show little ability to buffer the acidifying effects of elevated CO2 in their body compartments (Doney et al., 2009, Dupont et al., 2010b, Fabry et al., 2008, Melzner et al., 2009, Peck et al., 2009). Eurythermal species

40 however, have an enhanced ability at lower temperatures (Pörtner et al., 2000) but aerobic capacities are much lower in stenotherms (Portner and Sartoris, 1999) .

Ocean acidification effects at different latitudes

Tropical coral reefs calcify to form the massive three-dimensional structures that create habitat supporting extreme high biodiversity. As such, coral reef ecosystems are highly vulnerable to conditions that affect their ability to calcify or promote the dissolution of

CaCO3 (Hofmann et al., 2010). Simulations of climate models project that at a level of

550 ppm CO2, substantial dissolution of reefs worldwide is possible (Silverman et al., 2009)

Antarctic organisms may experience more variation than might be expected, which influences their resilience to future acidification (Hofmann et al., 2011). Living subarctic pteropods, Clio pyramidata, a major producer of aragonite, were exposed to a level of aragonite undersaturation projected for Southern Ocean surface waters by the year 2100, and shell dissolution occurred within 48 hours (Orr et al., 2005). It is hypothesised that polar pteropods, which only have one or two generations per year, will not be able to adapt quickly enough to live in the undersaturated conditions that will occur in high-latitude surface waters during the twenty-first century (Orr et al., 2005). Newly-settled subtropical juvenile clams (Mercenaria mercenaria) also showed shell dissolution and increased mortality when they were introduced to surface sediments that were undersaturated with respect to aragonite (a level that is typical of near shore, organic-rich surface sediments). Within two weeks of settlement, the shells were completely dissolved, leaving only the organic matrix (Green et al., 2004).

Although many physiological processes may be affected by rising ocean acidity, the declining aragonite and calcite saturation states of surface waters and establishment of corrosive conditions in some regions may particularly impact high-latitude planktonic and benthic calcifiers (Feely et al., 2009).

3.2 Aims

Because of these variations it is interesting to understand the degree of internal pH regulation in echinoderms from different climate regions. The main aim of this part of the study was to investigate the response of larval echinoderms to regulate their extracellular pH (pHe) in response to different environments, determining if there is inter-specific variation between latitudes when exposed to lowered seawater pHs. I hypothesise that polar species will be the most sensitive to pH change and show a delay in development, abnormal growth and have less ability to regulate pHe whereas the tropical species will show the smallest change in extracellular pH when exposed to acidified waters.

3.3 Study species: Evechinus chloroticus, Pseudechinus huttoni, Arachnoides placenta, Sterechinus neumayeri, and Odontaster validus

In this study, larvae from a range of echinoderm species; three regular echinoids (Evechinus chloroticus, Pseudechinus huttoni and Sterechinus neumayeri), one irregular echinoid (Arachnoides placenta) and one asteroid (Odontaster validus) were examined (Table 3.1, Figure 3.2)

Evechinus chloroticus (Valenciennes, 1846) is New Zealand’s most common endemic littoral and sub-littoral sea urchin inhabiting the entire coastline of New Zealand, as well as some southern and northern offshore islands (Dix 1970). It is commonly referred to as ‘Kina’ or ‘sea egg’ (McRae, 1959). Evechinus chloroticus is typically found in the sub-tidal region below 12m, but can be found in shallow intertidal waters in the North Island and can reach a maximum diameter of 19cm (Barker, 2001). This rocky bottom dweller is not often evenly distributed but found in clumps that can exceed densities of 50/m2 (Dix, 1970). Evechinus chloroticus has a broad diet but is typically algivorous, but if algae becomes sparse, it will scavenge on the substrate (Dix, 1970). Reproduction occurs annually during the summer months of December through to April (Lamare et al., 2002). Evechinus chloroticus larvae take about 4 days to reach the 4-rod pluteus stage (Clark, 2009). Adults are normally found in areas with moderate currents and wave action, avoiding highly exposed areas (Barker, 2007), with the

42 exception of Fiordland, where it is abundant in the deep fiords which indent the coastline (Barker, 2001).

Pseudechinus huttoni (Benham, 1908) is a small white urchin reaching a maximum diameter of 30 - 40 mm between the age of 6 - 11 years (Kirby et al., 2006). Pseudechinus huttoni is endemic to New Zealand, it inhabits the continental shelf in waters 30m to 500m deep (McClary and Sewell, 2003), and fjordic habitats in depths as shallow as 9m (Kirby et al., 2006). This rare fjordic emergence at these shallow depths is thought to reflect the deep-water-like environments of the New Zealand fiords, where low light, water motion, and sedimentation environments exist at relatively shallow depths (<40 m) (Kirby et al., 2006). Pseudechinus huttoni spawns in the winter months, with its annual breeding season from May through to September (Poorbagher et al., 2010), it is known to live on algae, yet it can be an omnivorous grazer, with contents of plants and coarse sediments recorded in its stomach (McKnight, 1969, Poorbagher et al., 2010).

Arachnoides placenta (Linnaeus, 1758) is a tropical sand dollar found throughout the Indo-Pacific region. It has a seasonal reproduction cycle, with gamete growth and accumulation occurring in December. Spawning occurs from March through to May coinciding with the decline in water temperature in March (26 - 28°C) (Haycock, 2004). Arachnoides placenta forms a dominant part of the intertidal fauna of many parts of North Queensland's coast. It plays an important role in reworking sediments and recycling nutrient’s in the benthic community of sandy shores (Bell and Frey, 1969). Larval development takes two weeks, during which they feed on microscopic algae (Chen and Chen, 1992). Arachnoides placenta larvae are very sensitive to high temperature, causing larval abnormalities (Chen and Chen, 1992). Larvae have also been shown to be sensitive to reduced seawater pH (Gonzalez-Bernat et al., 2013b) and temperature (Hardy et al., 2013). Decreases in developmental time to the pluteus stages was seen at warmer than ambient (24°C) temperatures, whereas cooler temperatures increased development. However, extreme cold (14°C) and warm (35°C - 37°C) temperatures led to abnormal development of larvae (Hardy et al., 2013).

Sterechinus neumayeri (Meissner, 1909) is the most abundant shallow-water Antarctic sea urchin, occurring on gravel rocks down to a depth of 500m (Bosch et al., 1987, Brey, 1991). It is endemic to Antarctica and has a circum-Antarctic distribution

(Brockington et al., 2001). Sterechinus neumayeri occurs in McMurdo Sound; it also exhibits slow growth (maximum size of 70 mm in 40 years) despite its relatively shallow distribution (< 20 m depth). Growth in this species is likely to be related to its polar distribution (Brey et al., 1995). Larvae reared at -1.8 to -0.9°C are capable of feeding 20 days after fertilisation and are competent to metamorphose after 115 days (Bosch et al., 1987). Spawning occurs during the late-austral spring to early summer months so that feeding larvae are in the plankton for the summer peak of phytoplankton abundance in McMurdo Sound (Bosch et al., 1987). Gametogensis is extensive and takes nearly 2 years to complete (Pearse et al., 1991). S. neumayeri has been described as a generalist feeder (McClintock, 1994).

Odontaster validus (Koehler, 1911) is a sea star widely distributed and abundant around the shallow benthic environments of Antarctica, most commonly distributed between 0 m and 200 m depth (Pearse, 1969), although there are reported cases of it being found at 940 m (Dearborn, 1977). Odontaster validus is an opportunist feeder, acting as a scavenger, suspension feeder, herbivore and active predator (Pearse, 1964). Odontaster validus has been said to act as a keystone species as it feeds on benthic larvae, which can in turn influence community structures on the Antarctic shelf and regulate populations of space dominating sponges, other seastars and nudibranchs (Dayton et al., 1974, McClintock et al., 1988). O. validus spawns in the austral winter (late May - mid September). Gametogenisis can begin up to 2 years before spawning (Pearse, 1969). Feeding larvae develop in the water column for 6 month before settling (McClintock and Pearse, 1986), which ensures widespread dispersal of larvae (Pearse et al., 1986).

Table 3.1 Details on the collection location, experimental conditions and reproductive period of the five study species: Evechinus chloroticus, Pseudechinus huttoni, Arachnoides placenta, Sterechinus neumayeri and Odontaster validus.

Species E. chloroticus P. huttoni A. placenta S. neumayeri O. validus Collection Site Doubtful Sound, Doubtful Sound, Pallaranda, Cape Evans, Cape Evans, Fiordland, Fiordland, Townsville, Antarctica Antarctica New Zealand New Zealand Australia 77°57’S, 77°57’S, 166°44’E Latitude 42°25’S, 42°25’S, 19°12’04’’S, 166°44’E Longitude 170°56’E 170°56’E 146°46’24”E Number of specimens collected 30 30 30 20 20 Ambient pH at collection site 8.16 8.16 8.08 8.01 8.01 Ambient temperature at 15 13 24 -1.9 -1.9 collection site (°C) Collection depth (m) 12-15m 15-25m Sand/low tide 10-20m 10-20m Habitat Rocky Surfaces Rocky surfaces Sandy sub-tidal Mud, rock, sand Mud, rock, sand Spawning Period Dec-Apr(a) May-Sept(b) April- May(c) Sept-Dec(d) May-Sept(d) Experimental data Experimental pH(nbs) 8.1, 7.8, 7.6, 7.4 8.1, 7.8, 7.6, 7.4 8.08, 7.8, 7.6, 7.4 8.01, 7.7, 7.5, 7.2 8.01, 7.7, 7.5, 7.2 Experimental temperatures (°C) 14 ± 1 10 ± 1 25 ± 1 -1.9 -1 ± 1 Experimental Period Dec-Jan Aug May Oct Jun-Jul Number of specimens spawned 8 6 6 10 7

(a) (Lamare et al., 2002) (b) (Poorbagher et al., 2010) (c) (Gonzalez-Bernat et al., 2013b) (d) (Chiantore et al., 2002)

a. Evechinus chloroticus b. Arachnoides placenta

c. Pseudechinus huttoni d. Sterechinus neumayeri

e. Odontaster validus

Figure 3.2 Adult echinoderms used in this study: a. Evechinus chloroticus b. Arachnoides placenta, c. Pseudechinus huttoni, d. Sterechinus neumayeri, e. Odontaster validus. Scale bars represent 10mm.

3.4 Materials and methods

3.4.1 Animal collections

Adult Evechinus chloroticus and Pseudechinus huttoni were collected on 29 January and 3 July 2012 respectively by SCUBA from Doubtful Sound, Fiordland (Figure 3.3 A). Animals were kept in a flow-through system at the Portobello Marine Lab in 14 ± 1°C (E. chloroticus) and 10 ± 1°C (P. huttoni). Adult Arachnoides placenta were collected from the low tide mark at the sandy Pallarenda beach, Townsville, Australia on 24 May 2012 (during their natural spawning month) (Figure 3.3 B) and were kept in a flow through system at 25 ± 1°C (Table 3.1). Odontaster validus were available in the Portobello Marine Lab from prior collection from Antarctica in 2009 where they were air transported to New Zealand and maintained in a controlled temperature cabinet of -0.5°C and fed the bivalve (Austrovenus stutchburyi) flesh monthly. Sterechinus neumayeri were collected on the 27 October 2012 by remote operated vehicle (ROV) though a dive hole situated at Cape Evans, McMurdo Sound, Antarctica (Figure 3.3 C). Specimens were kept in a flow-though tank at -1.91°C.

a Doubtful Sound

Lat 42°16’50"S, Long 166°51’E 10km

Townsville Lat 19°12’04’’S, Long 146°46’24”E

200km

Townsville

Cape Evans Lat 77°57’18"S, Long 166°44’25"E 15km

Figure 3.3 Location of (a). Doubtful Sound, Fiordland, New Zealand, where Evechinus chloroticus and Pseudechinus huttoni were collected (b). Palleranda Beach, where tropical sand dollar Arachnoides placenta was collected and (c). Cape Evans, Antarctica, where Sterechinus neumayeri and Odontaster validus were collected (McClintock et al., 1988, Tremblay, 2013) . 48

A C 3.4.2 Spawning and culturing Australia AntarcticaNew Zealand B Spawning and rearing of E. chloroticus and P. huttoni was conducted at the Portobello Marine laboratory, A. placenta at AIMS, Australia and S. neumayeri at Scott Base, Antarctica. Spawning was induced by an inter-coelomic injection of 0.5 mol/l of KCl using standard techniques (Lamare et al., 2006) and gametes were collected in 500 ml beakers of filtered seawater (1µm) with temperatures maintained using a temperature controlled room (Table 3.1, Figure 3.4). Urchins were left to spawn for 30 min or until spawning had ceased. Depending on species, three males and five to seven female urchins were spawned, eggs and sperm were pooled for each species to avoid maternal and paternal effects. Eggs were cleaned by gently washing three times in a 200 ml beaker of fresh seawater. Two drops of concentrated sperm were then added to the 200 mls and gently stirred to achieve fertilisation. Only batches with >95 % fertilisation were used. To avoid the effects of pH on fertilisation, this was carried out in ambient seawater. Following fertilisation, embryos were washed three times in filtered seawater before being transferred to 50 ml falcon tubes fitted with four 100 µm mesh windows (5 x 0.5 cm) in the side to allow for water flow (Figure 3.5).

Tubes were placed in a holder in a flow through system of filtered seawater maintained at experimental pH levels of 7.8, 7.6, 7.4 and ambient (pH 8.1) in a temperature controlled room of 18°C (E. chloroticus and P. huttoni) and 25°C for A. placenta. For S. neumayeri embryos and larvae were raised in pH 7.7, 7.5, 7.2 and ambient (pH 8) at - 1.9°C. Embryos were raised in this system at a concentration of ~10 ind/ml for 1 - 21 days, depending on when they had reached the 4-armed pluteus stage. Throughout the experiment E. chloroticus, P. huttoni and S. neumayeri were fed (once larvae had reached the feeding stage) 2 ml of the phytoplankton Isocrysis and Dunaliella spp. at a concentration of 4000 – 6000 cells/ml every two days. The faster growing A. placenta larvae were not fed due to its short development time (24 hour pluteus).

Odontaster validus spawning, conducted at the Portobello Marine laboratory, was induced by intercolemetric injection of 0.5 ml per individual of the ovulatory hormone 1-methyladenine (1MA) (10-3 to 10-5 M concentrations). Five females were spawned and two males were dissected as they did not spawn with the hormone. Eggs were

49 fertilised by adding three drops of sperm from the freshly dissected males to beakers of filtered seawater 30 mins after spawning. A fertilisation rate of 90-100% was observed. Embryos were washed three times and transferred to 500 ml containers, at pH 7.7, 7.5, 7.2 and ambient (pH 8). Larvae were fed 1 ml of Danelliella (4000 – 6000 cells/ml ) per 500 ml container once a week when they had reached the feeding stage (2 week after fertilisation).

Figure 3.4 Spawning female Sterechinus neumayeri (left) and male Pseudechinus huttoni (right) after being injected with KCL to induce spawning.

Window (5 x 0.5cm) Falcon Tube Mesh (100µm)

Figure 3.5 50ml falcon tube used to rear larvae in flow-through seawater systems. Four, 5 x 0.5cm windows were made equally around tube, covered by 100 µm mesh to retain larvae but to allow water to flow through. 50Scale bar represents 1cm.

3.4.3 Adjustment and monitoring of seawater parameters

Due to logistical considerations the setup for changing seawater pH for each species varied. For each experiment four treatments were applied, three of which approximated projected ρCO2 levels for the next 300 years of pH: 7.8, 7.6 and 7.4, as well as an ambient treatment (for tropical and temperate experiments) (Table 3.2). For both polar experiments the lowered pH was 7.7, 7.5 and 7.2 due to Antarctic water having a naturally lower initial seawater ρCO2.

Carbonate chemistry

In all treatments, pH was regulated to ± 0.04 units of the desired pHnbs (all pH on NBS scale) using a tunze pH/CO2 controller to control pH (7074/2 TUNZE AQUA RIENTECHNIK GMBH, Pennsbury Germany), which had been calibrated to buffers 4.0, 7.0 and 9.0 at 20oC (Pronalys biolab). Independent measurements were taken daily using a Mettler Toledo MP220 pH meter, to ensure pH was consistently ± 0.04 of the regulator readings. A seawater sample was also taken and fixed in mercuric chloride for later carbonate analysis. The carbonate chemistry of ambient and treated seawater was calculated using measured pH and total alkalinity (AT). Both pH and AT values were used as input parameters in the program SWCO2 (Hunter 2009) to determine partial pressure of CO2 and calcite and aragonite saturation states.

Water samples from each collection site and the flow-through systems were fixed with saturated mercuric chloride (HgCl2) for later laboratory alkalinity analysis and dissolved ρCO2, using a closed cell potentiometric titration and analysed by least squares linear optimisation technique. A quality control was performed based on the

CO2 Certified Reference Material supplied by AG Dickson, Scripps Institution of

Oceanography. Partial pressure of CO2 (ρCO2) and the saturation values for calcite

(ΩC) and aragonite (ΩA) were calculated using the computer program SWCO2 (Hunter

2009). Seawater properties were determined using the CO2 equilibrium constants given by Mehrbach et al. (1973).

Evechinus and Pseudechinus

A flow-through system was set up in a controlled temperature room of 18°C. 100%

CO2 gas was bubbled into 3 separate filtered seawater 70 L header tanks with the water level controlled by a ball float and fitted with a 130µm TAGLINE screen filter from the seawater intake, a water pump to mix water and tunze pH/CO2 controller to control pH (7074/2 TUNZE AQUA RIENTECHNIK GMBH, Pennsbury Germany). Each header tank fed three smaller 12 L replicate trays (50 cm x 30 cm x 8 cm) with flowing seawater continuously at a rate of 1 L/min (Figure 3.6). The consistent flowing water system meant dissolved oxygen concentrations were maintained at ambient. Four falcon tubes were placed in each tank and secured to a tray that keep them submerged (Figure 3.6 and 3.7 a). For the summer E. chloroticus experiment water temperature was 15°C, with water at 10°C for P. huttoni in the winter.

Figure 3.6 pH controlled flow-through larval rearing system for New Zealand experiments with 1, 2 and 3 representing the replicate tanks.

Arachnoides placenta

A flow-through was set up in a controlled temperature room of 25°C. Limited equipment meant that I could only set up one pH treatment at a time. 100% CO2 gas was bubbled into a 50 L tank with a water pump to mix water and tunze pH/CO2 controller (7074/2 TUNZE AQUA RIENTECHNIK GMBH, Pennsbury Germany). This tank fed into a smaller 40 L tank at 2 L/min and five falcon tubes were placed and secured to a tray that keep them submerged (Figure 3.7 b). Seawater temperature was 25°C.

Sterechinus neumayeri

A flow-though was set up using water with a constant temperature of -1.9°C. 100%

CO2 was bubbled into a 70 L tank with a water pump to mix water and monitored and regulated CO2 by a tunze pH/CO2 controller (7074/2 TUNZE AQUA RIENTECHNIK GMBH, Pennsbury Germany). The 70 L tank was set for pH 7.5, this fed into a 50 L tank below so the pH 7.5 was well mixed, this water then fed into another 50 L tank that had a fresh seawater input at a rate of 1L/min to result in a pH of 7.7, fitted with a water mixer and closely monitored. Five falcon tubes were fitted into trays and submerged in both the pH 7.7 and pH 7.5 treatment (Figure 3.7 c). A 7.2 treatment was made by bubbling 100% CO2 into a 20 L bucket until the desired pH was reached, larvae for this treatment were reared in closed 500 ml plastic jars at 10 ind/ml and water was changed every 5 days to prevent pH and oxygen changes. Seawater temperature was -1.9°C .

Odontaster validus

Due to physical containment level 2 (PC2) laboratory restrictions an open flow-through system was not able to be constructed, therefore O. validus embryos were placed in 500 ml plastic jars of seawater at pH treatments 7.7, 7.5, and 7.0 using methods previously applied to the species (Gonzalez et al. 2013). This was accomplished by bubbling CO2 into 10 L buckets containing filtered seawater and checked with a (Mettler toledo MP220) pH meter calibrated with NBS buffers 4.0, 7.0 and 9.0 at 20°C (Proalys biolab). Seawater pH was monitored until the desired pH was reached and stable (±

0.04 pH units). Embryos were added to the jars at a concentration of 10 ind/ml, then sealed with no air spaces to prevent equalisation with the atmosphere. Jars were kept in a constant refrigerated temperature of -1°C. There were three replicates for each pH treatment. To keep oxygen and pH levels optimal, seawater in jars was changed once a week using 100µm mesh and a larval cleaning tube.

Larval rearing tubes clipped 12 L Replication trays into submerged tray a

b c

Figure 3.7 Photos of flow through systems for each species: a. E. chloroticus/P.

huttoni, b. A placenta, and c. S. neumayeri

3.4.4 Morphometrics

To determine morphometric differences in early cellular development of larvae, ambient light photographs were analysed to record the percentage of embryos which were abnormal in each pH treatment. Embryos that were not spherical and had irregular sized blastomeres were considered abnormal (Table 3.3, Appendix 1). To determine changes in the pluteus stage, four body components were established as proxies to identify differences in morphology from experimental conditions. The measured body components were: body length (BL), body width (BW), post oral rod length 1 (R1), and post oral rod length 2 (R2) (Figure 3.8). For gastrula and sea star bipinnaria only length and width were measured.

Ten larvae from each replicate pH treatment were photographed and measured during several stages of development. For E. chloroticus and P. huttoni, images were taken on days 1 and 12; A. placenta on days 1 and 3; S. neumayeri on days 1, 7 and 28, and; O validus on days 1, 15 and 43 (Table 3.3). All larval measurements were made using Image J software Version 1.42q.

a TL

R2 R1

TL TL

Figure 3.8 Morphometric measurements of a) sea urchin larvae, using Evechinus chloroticus as an example and b) Odontaster validus gastrula and bipinnaria. Measurements made are total length (TL), total width (TW), and post-oral skeletal arms 1 and 2 (R1, R2) on pluteus larvae. Pluteus larvae were also viewed under normal and polarised light to highlight the larval skeleton. Scale bar represents 100 µm.

3.4.5 Extracellular pH measurements

Using methods described in 2.23, a standard curve was made for each species (Appendix 2). During several stages of development (days indicated above) a 5 ml sample of water containing embryos/larvae was taken from each pH treatment and transferred to a 50 ml falcon tube that contained a 30 µM solution of the fluorescent probe HPTS (8-hydroxypyrene-1,3,6-trisulfonic acid) adjusted to the equivalent treatment pH conditions. During incubation, HPTS was taken up by the larvae into the extracellular space. In most cases embryos and larvae were left for a twelve hour incubation period in the probe, however for the faster growing A. placenta incubation was four hours to allow imaging of earlier cellular development.

After incubation, larvae were washed four times in a Bogerov counting chamber of filtered seawater (of the same pH) to remove of any excess HPTS in the surrounding water. Ten larvae from each pH treatment were imaged on a slide in a dark room using a Nikon fluorescent microscope (Eclipse TE300) fitted with a Nikon filter cube (exciters UV1A and B2A, band pass filter HQ535/50) and plan fluor dry objectives (20x, 40x, and 100x). Exposure time was set at 2 seconds (see chapter 2) (Figure 3.9).

Arachnoides placenta was photographed using a ZEISS AxioCam MRc5 camera under excitations 358 nm (DAPHI) and 495 nm (FITC), and emission maximum of 461 nm and 525 nm respectively. This wavelength covered the absorbance maxima required for HPTS. Images were photographed using the imaging Axiovision Rel 4.8 Application software. Exposure time was set at two seconds.

Visible light

Oral hood

Gut Oesophagus

405nm 495nm

Figure 3.9 Images of larvae after being incubated in HPTS, washed, excited at wavelengths 405 nm/495 nm and photographed at emission 520 nm (Odontaster validus as an example). Scale bar represents 100 µm.

Due to the movement of larvae on the slide under the microscope only the slow moving Antarctic species were able to be photographed at the blastula and gastrula stage. These larvae remained still enough to allow the 405/490nm images to line up (for two consecutive two second exposure photos). Multiple methods were attempted to limit movement of other species such as using a glycerine based gel to slow larvae but this method affected the wellbeing of the larvae, therefore blastula and gastrula were excluded from other treatments.

3.4.6 Statistical analysis

Morphometrics

A one-way ANOVA was used to identify significant differences among body component lengths of larvae reared under different seawater pH treatments (P < 0.05). The data set consisted of 40 observations for each species and two variables for early embryo development (abnormal and normal). Two variables were used for gastrula and bipinnaria (body length (BL) and body width (BW)) and 4 variables for pluteus stages (body length (BL), body width (BW), post oral rod 1(R1), post oral rod 2 (R2)). A Turkey's HSD post-hoc test was carried out to identify significant differences among pH treatments (P < 0.05). Multiple t-test (α = 0.05) were run on the post oral rods to distinguish any significant asymmetry between R1 and R2 (p < 0.05). All analyses were generated using JMP® 7.0 (2007, SAS Institute Inc., USA). pHe

Fluorescent imaging processing of larvae was carried out using the techniques described in chapter 2.2.2. The species-specific standard curve equation was used to transform the resulting ratio into a colour map of extracellular pH. Photographs where the 460/405nm image overlap did not exactly line up due to larvae moving were excluded, leaving some sample sizes less than ten.

Regional variations in extracellular pH, (for example larval arms, oesophagus, gut) (see Figure 2.6) were calculated by drawing a polygon on the pH map in MATLAB which

58 would generate the average pHe from within the selected area. From each of the four treatments, pHe measurements were made from ten larvae. All pHe measurements were log transformed prior to statistical analysis. The average pH ratios from each treatment in each region was analysed using a one-way ANOVA (P < 0.05) with Turkey's HSD post-hoc test to identify significant differences among pH treatments (P < 0.05). ANOVA assumptions were examined through a visual examination of the Normal Quantile Plots of the standard deviation (σ) and a Bartlett test confirmed homogeneity of variance. Heterogeneous variances were compared using Welch ANOVA testing for equal means, allowing for unequal variances. Statistical analyses were carried out using JMP® 7.0 (2007, SAS Institute Inc., USA).

3.5 Results

3.5.1 Adjustment and monitoring of seawater parameters

For Evechinus chloroticus the ρCO2 was 440 µatm under ambient conditions. For treated seawater, the ρCO2 approximately doubled in pH 7.8 (989 µatm), increased 4- fold in pH 7.6 (1636 µatm) and ~ 6-fold in pH 7.4 (2614 µatm). Calcite and aragonite were saturated under ambient conditions (ΩC = 3.7 and ΩA = 2.4, respectively) and calcite for pH 7.8 (ΩC = 1.8) while aragonite was almost undersaturated for pH 7.8 (ΩA

= 1.1). For pH 7.6 calcite remained saturated while aragonite was undersaturated (ΩC =

1.2 and ΩA = 0.75). In pH 7.4, both calcite and aragonite were under saturated (ΩC =

0.74 and ΩA = 0.48) (Table 3.2 a).

For Pseudechinus huttoni ρCO2 was 383 µatm under ambient conditions. For treated seawater, a 2.6-fold increase was recorded in pH 7.8 (983 µatm), 4-fold increase in pH 7.6 (1605 µatm) and almost 7-fold increase in pH 7.4 (2592 µatm). Calcite and aragonite were saturated under ambient conditions (ΩC = 3.6 and ΩA = 2.3) and for pH

7.8 (ΩC = 1.7, ΩA=1.1) For pH 7.6 calcite was almost under saturated (ΩC = 1.1) and aragonite was undersaturated (ΩA = 0.75) and pH 7.4 (ΩC = 0.70 and ΩA = 0.45) both calcite and aragonite were under saturated (Table 3.2 b).

For Arachnoides placenta ρCO2 was 529 µatm under ambient conditions. For treated seawater, a 1.4-fold increase was recorded in pH 7.8 (716 µatm), 1.6-fold increase in pH 7.6 (845 µatm) and a 3.4-fold increase in pH 7.4 (1785 µatm). Calcite and aragonite were saturated under ambient conditions (ΩC = 4 and ΩA = 2.8), pH 7.8 (ΩC = 3.3, ΩA =

2.2) and pH 7.6 (ΩC = 2.9, ΩA = 1.9). In treatment pH 7.4 calcite was still saturated (ΩC

= 1.63) and aragonite was almost undersaturated (ΩA = 1) (Table 3.2 c).

For Sterechinus neumayeri ρCO2 was 413 atm under ambient conditions. For treated seawater, a 3.3-fold increase was recorded in pH 7.7 (1344 µatm), 4.4-fold increase in pH 7.5 (1773 µatm) and a 8.7-fold increase in pH 7.2 (3710 µatm). Calcite and aragonite were saturated under ambient conditions (ΩC = 2.1 and ΩA = 1.3), from pH

7.7 calcite and aragonite were both undersaturated (ΩC = 0.70, ΩA = 0.44), pH 7.5 (ΩC

= 0.55 and ΩA = 0.35) and pH 7.2 (ΩC = 0.27, ΩA = 0.17) (Table 3.2 d).

For Odontaster validus ρCO2 was 407 atm under ambient conditions. For treated seawater, a 2.5-fold increase was recorded in pH 7.7 (1018 µatm), 4.3-fold increase in pH 7.5 (1744 µatm) and a 9-fold increase in pH 7.2 (3526 µatm). Calcite and aragonite were saturated under ambient conditions (ΩC = 2 and ΩA = 1.6), for all other treatments calcite and aragonite were both undersaturated (pH 7.7; ΩC = 0.84, ΩA = 0.53, pH 7.5;

ΩC = 0.54, ΩA = 0.34 and pH 7.2; ΩC = 0.28, ΩA = 0.17) (Table 3.2 e).

Table 3.2 Temperature (°C), salinity (PSU), pH (NBS scale), total alkalinity (µmol -1 1 kg soln- ), partial pressure of CO2 (ρCO2), CaCO3 saturation value for calcite (ΩC)

and aragonite (ΩA) for ambient and treated seawater. (a) Evechinus chloroticus (b) Pseudechinus huttoni (c) Arachnoides placenta (d) Odontaster validus (e) Sterechinus neumayeri.

(a) Evechinus chloroticus, Otago Harbour, Ambient T =15°C (±1), Ambient salinity = 35 (±1)

pH NBS 8.16 7.8 7.6 7.4

Alkalinity(AT) 2241.1 2241 2268 2241

ρCO2 (µatm) 440 989 1636 2614

ΩCalcite 3.7 1.8 1.2 0.74

Ωaragonite 2.4 1.1 0.75 0.48 (b) Pseudechinus huttoni, Otago Harbour, Ambient T = 10°C(±1), Ambient salinity = 35(±1)

pH NBS 8.16 7.8 7.6 7.4

Alkalinity(AT) 2275.4 2270 2270 2270

ρCO2 (µatm) 383 983 1605 2592

ΩCalcite 3.6 1.7 1.1 0.70

Ωaragonite 2.3 1.1 0.70 0.45 (c) Arachnoides placenta, Townsville, Ambient T =25°C, Ambient salinity = 35(±1)

pH NBS 8.08 7.8 7.6 7.4

Alkalinity(AT) 2283 2284 2288 2288

ρCO2 (µatm) 529 716 845 1785

ΩCalcite 4 3.3 2.9 1.63

Ωaragonite 2.8 2.2 1.9 1

(d) Sterechinus neumayeri, McMurdo Sound, Ambient T = -1.9°C, Ambient salinity = 34.82

pH NBS 8.01 7.7 7.5 7.2

Alkalinity(AT) 2351 2328 2345 2346

ρCO2 (µatm) 413 1344 1773 3710

ΩCalcite 2.1 0.70 0.55 0.27

Ωaragonite 1.3 0.44 0.35 0.17

(e) Odontaster validus, Otago Harbour, Ambient T = -1°C, Ambient salinity = 35(±1)

pH NBS 8.01 7.7 7.5 7.2

Alkalinity(AT) 2275 2268 2270 2275

ρCO2 (µatm) 407 1081 1744 3526

ΩCalcite 2.0 0.84 0.54 0.28

Ωaragonite 1.6 0.53 0.34 0.17

3.5.2 Morphological observations of early cell development

In all species, the overall trend showed either delay in development or increasing percentage of cell abnormality with decreasing seawater pH, characterised by less spherical cells and an increasing percentage of ruptured cell walls (Figure 3.10, 3.11 and Appendix 1, 3). P. huttoni showed the greatest percentage of abnormal embryos

(80%) in the lowest pH treatment (pH 7.4) (F(3,31) = 59.8, P < 0.0001). Sterechinus neumayeri showed significantly abnormal cell development in treatments 7.5 and 7.2 with both treatments consisting of 54% abnormality (F(3,26) = 5.3 P < 0.05) (Figure 3.10, 3.11 and Appendix 2,3). E. chloroticus and O. validus showed no significant abnormalities in the first cellular stage. Significant abnormalities in cell development was, however, noticed in E. chloroticus 2-cell and 4-cell stage with 80 - 90% abnormal embryos in treatment pH 7.6 and pH

7.4. (F(3,39) = 93.7, p < 0.0001). Pseudechinus huttoni experienced a delayed development with decreasing seawater pH (Figure 3.10, Appendix 1), resulting in 100% and 97% developed to the four-cell stage abnormally in treatments pH 7.6 and pH 7.4 respectivly (Figure 3.11 and Appendix 3). For O. validus 20% of embryos showed blastoderm cell wall irregularity and an increasing number of bumps on the outer cell wall when raised under pH 7.2, yet this was not significantly different from ambient which consisted of 6.7% abnormality. For A. placenta blastula only the lowest pH 7.4 treatment had a significantly higher percentage of abnormal embryos (F(3,23) = 25.3 P < 0.001) (Figure 3.10, 3.11 and Appendix 2, 3).

Ambient pH (8.1) pH 7.8 pH 7.6 pH 7.4

Fertilisation Blastomere Evechinus chloroticus (Day 1) envelope

Pseudechinus huttoni (Day 1)

Egg envelope Arachnoides placenta (Day 1) Blastderm Blastocoel

Sterechinus neumayeri (Day 1) Blastomeres Ambient pH (8.0) pH 7.7 pH 7.5 pH 7.2

Odontaster validus (Day 5) Fertilisation envelope Blastoderm Blastocoel

Figure 3.10 Early embryological development of E. chloroticus, P. huttoni, A. placenta, S. neumayeri and O. validus when raised under four seawater pH treatments. Scale bars represent 50µm.

a) Abnormal early cell development (1 cell stage)

Evechinus chloroticus Pseudechinus huttoni Sterechinus neumayeri 100 100 c 100 80 80 80 b 60 b 60 60 40 ab b 40 ab 40 20 a a 0 20 20 Perccentage Abnormal SE) (± Perccentage Percentage abnormal (± SE) abnormal (± Percentage

0 SE) abnormal (± Percentage 0 Ambient 7.8 7.6 7.4 Ambient 7.7 7.5 7.2

b) Abnormal 2-cell stage

Odontaster validus Evechinus chloroticus Pseudechinus huttoni 100 100 100 b b 80 80 80

60 c 60 60 40 40 a a 40 b 20 20 20 a Percentage abnormal (± SE) abnormal (± Percentage Percentage abnormal (± SE) abnormal (± Percentage 0 SE) abnormal(± Percentage 0 0 Ambient 7.7 7.5 7.2 Ambient 7.8 7.6 7.4 Ambient 7.8 7.6 7.4

C) Abnormal 4-cell stage d) Abnormal blastula

Evechinus chloroticus Pseudechinus huttoni Arachnoides placenta 100 b b b b 100 100 90 b 80 80 70 80 60 a b 60 60 50 40 40 40 30 a a a 20 20 20 a

Percentage abnormal (± SE) abnormal (± Percentage 10 Percentage abnormal (± SE) abnormal (± Percentage Percentage abnormal (± SE) abnormal (± Percentage 0 0 0 Ambient 7.8 7.6 7.4 Ambient 7.8 7.6 7.4 Ambient 7.8 7.6 7.4

pH treatment

Figure 3.11 Percentage of a) Abnormal early egg development, b) Abnormal 2-cell embryonic stage c) Abnormal 4-cell embryonic stage and d) Abnormal blastula, of five echinoderm species when larvae were grown in either ambient pH seawater, or lowered pH seawater3.5.2 pH.Extracellular Different letters pH indicates significant differences between pH treatments (P<0.05) (N = 30).

3.5.3 Morphological observations of Larvae - gastrula, pluteus and sea star bipinnaria developmental stages

For E. chloroticus there was a significantly narrower body width between treatment pH

7.8 (408 µm) and pH 7.4 (348 µm) (Width: F(3, 39) = 3.12 P=0.04) (Figure 3.13) . Post oral rod 1 measured 335 µm under pH 7.8 which was significantly shorter under pH

7.4 at 290 µm (Rod1: F(3, 39) = 3.83, P < 0.05). Post oral rod 2 ranged from 327 µm to 340 µm in treatments 7.6, 7.8 and ambient pH and in treatment 7.4 length was significantly shorter at 284 µm (Rod2: F(3, 39) = 5.3, P < 0.01) (Figure 3.12, Appendix 4). Ambient pH length was 415µm which was similar to the lowest pH treatment of a length of 413 µm.

Pseudechinus huttoni had a number of significant differences among morphometric measurements, with the pH 7.4 treatment being significantly shorter in all body measurements. Mean body length in ambient pH was 446 µm, which was significantly shorter (381 µm) under the lowest pH treatment (BL: F(2, 29) = 16.47, P < 0.0001). Mean pluteus width under ambient pH was 294 µm which significantly decreased to 228 µm in treatment pH 7.4. BW: F(2, 29) = 9.41, P < 0.001). Under ambient pH post oral rods 1 and 2 were 230 µm and 223 µm respectively which significantly decreased to 171 µm and 179 µm respectively (R1: F(2, 29) = 15.38, P < 0.0001, R2: F(2, 29) = 10.58, P < 0.001). (Figure 3.12, Appendix 4).

Arachnoides placenta had significantly shorter lengths in all morphometric measurements under treatment pH 7.4. Ambient pH length was 357 µm which was significantly shorter (304 µm) under pH 7.4 (F(3, 39) = 17.65, P < 0.001). Pluteus width significantly decreased from an ambient width of 329 µm to 291 µm and 278 µm in treatments 7.6 and 7.4, respectively (F(3, 39) = 7.4, P < 0.001). Ambient rod length for R1 and R2 was 299 µm and 303 µm, respectively, which was significantly shorter under treatment pH 7.4 at 243 µm and 244 µm respectively (R1: (F(3, 39) = 6.76, P <

0.001, R2: F(3,39) = 6.73, P < 0.001)). (Figure 3.12, Appendix 4).

Sterechinus neumayeri gastrula showed a significantly shorter length from 151 µm under ambient pH conditions to 137 µm under pH 7.5 (F(3, 39) = 20.51, P < 0.001),

65 whilst width was not affected by a lowering of pH (Figure 3.12, Appendix 4). The four week old pluteus width was significantly greater in the pH 7.2 treatment measuring 123

µm than in ambient (103 µm) (F(3, 39) = 6.66, P < 0.001). Length was not affected by pH, and only rod 1 was significantly shorter in pH 7.4 than treatment 7.8 by 10 µm (Figure 3.12, Appendix 4).

Similar trends were observed in O. validus where the gastrula width was also significantly wider in pH 7.2 (270 µm) than in ambient pH (235 µm) (F(3, 39) = 7.31, P < 0.001), while length was not significantly different (Figure 3.12, Appendix 4). The opposite was seen in the 43-day old bipinnaria, average ambient length was 728 µm which significantly shorter in the 7.2 treatment at 607 µm (F(3, 39) = 20.51, P < 0.001). Ambient width was 417 µm which significantly narrower (304 µm) in the pH 7.4 treatment (F(3, 39) = 21.75, P < 0.001) (Figure 3.12, Appendix 4).

There was no asymmetrical differences between post-oral rods 1 and 2 in any of the echinoid species (Figure 3.13, Table 3.3)

a) Evechinus chloroticus (n=30) b) Arachnoides placenta (n=30) 600 600

500 500 ab a ab Ambient a Ambient 400 a a 400 b b a a ab a ab a a a a a 7.8 SE m)± a a a a m)± SE m)± b 7.8 b µ b

µ b 300 300 b b 7.6 7.6 200 200

7.4 Length (

Length ( 7.4 100 100

0 0 Length Width Rod1 Rod2 Length Width Rod1 Rod2

d) Sterechinus neumayeri gastrula (n=30) c) Psudechinus huttoni (n=30) 600 600 500 a a 500 b Ambient 400 Ambient 400 7.7 m)± SE m)± a a SE m)± µ 300 7.8 µ 300 b a a a a 7.5 b 200 b 7.4 200 a a ab b 7.2 Length ( Length ( 100 100

0 0 Length Width Rod1 Rod2 Length Width e) Sterechinus neumaeryi 4 weeks (n=30) f) Odontaster validus gastrula (n=30) 600 600

500 500

Ambient 400 400 Ambient m)± SE m)±

m)± SE m)± 7.7 µ c 7.7

µ 300 300 a a 7.5 b 7.5 200 200 Length (

Length ( 7.2 ab b 7.2 a a ab 100 aab b 100 0 0 Length Width Rod1 Rod2 Length Width

g) Odontaster validus bipinnaria (n=30) 800 a 700 bc b c 600 Ambient 500 a a a

SE m)± µ 7.7 400 b 300 7.5 Length ( 200 7.2 100

0 Length Width Morphometric Measurment

Figure 3.12 Length of morphometric variables in larvae of five species grown in either ambient pH seawater, or lowered pH seawater a) Evechinus chloroticus 12 day old pluteus b) Arachnoides placenta four day old pluteus, c) Pseudechinus huttoni 12 day old pluteus, d, e) Sterechinus neumayeri four week old pluteus and f, g) Odontaster validus 43 day old bipinnaria. While similar in developmental stage (4-armed pluteus or gastrula), larvae varied in age. Columns having different lower case67 indicat es significant differences among pH treatments (P<0.05) (N = 30).

a Evechinus chloroticus b Arachnoides placenta 400

350 400 300 350 250 300 m) ± SE m) ± 250 m) ± SE m) ± µ 200 POA1 POA1 µ 200 150 POA2 150 POA2 100 100 Length ( 50 50 Length ( 0 0 Ambient 7.8 7.6 7.4 Ambient 7.8 7.6 7.4 Seawater pH Seawater pH

c Pseudechinus huttoni d Sterechinus neumayeri

400 400

350 350 300 300 250 250 m) ± SE m) ± m) ± SE m) ± µ µ 200 POA1 200 POA1 150 150 POA1 100 100 POA2 Length ( Length ( 50 50 0 0 Ambient 7.8 7.4 Ambient 7.8 7.6 7.4 Seawater pH Seawater pH

Figure 3.13 Mean length of post oral arms 1 and 2 for a) E. chloroticus b) A. placenta c) P. huttoni and d) S. neumayeri under ambient and three reduced seawater pH levels. n =10. Error bars are ± standard error.

Table 3.3 Students t-test to identify significant asymmetrical differences between the post oral arm 1 and post oral arm 2 on the pluteus from a) E. chloroticus b) A. placenta c) P. huttoni , d) S. neumayeri . n=10. Welch t-testing for equal means, allowing for unequal variances. a E. chloroticus b A. placenta Ambient t = 2.1, p > 0.05, df = 17.8 t = 2.1, p > 0.05, df = 16.0 7.8 t = 2.1, p > 0.05, df = 17.8 t = 2.1, p > 0.05, df = 17.0 7.6 t = 2.1, p > 0.05, df = 17.9 t = 2.1, p > 0.05, df = 15.1 7.4 t = 2.1, p > 0.05, df = 16.5 t = 2.1, p > 0.05, df = 17.6 c P. huttoni d S. neumayeri Ambient t = 2.1, p > 0.05, df = 17.9 t = 2.1, p > 0.05, df = 18.0 7.8 t = 2.1, p > 0.05, df = 15.5 t = 2.1, p > 0.05, df = 18.0 7.6 n/a t = 2.1, p > 0.05, df = 11.2 7.4 t = 2.1, p > 0.05, df = 16.3 t = 2.1, p > 0.05, df = 17.7

3.5.4 Extracellular pH (pHe) of Larvae - gastrula, pluteus and bipinnaria developmental stages

Extracellular pH in E. chloroticus gut varied between pHe 6 and pHe 8.02 across pH treatments with a mean of 7.9 under ambient (n = 8 SE = 0.1). Gut pHe significantly decreased among pH treatments and as seawater pH was lowered (F(3,31) = 34.56, P<0.001). Treatment pH 7.8 was significantly lower than ambient and 7.6 and 7.4 were significantly lower than ambient and 7.8. Mean pHe in the gut wall varied between pH 5.95 and 7.17, with a mean of pHe 6.9 ( n = 8, SE = 0.08) under ambient conditions.

This significantly decreased in treatments 7.6 and 7.4 (F(3, 31) = 18.8, P<0.0001) (Figure 3.14 & 3.15, Table 3.4). In the oesophagus and larval arms regions there was a significant difference among pH treatments and pHe (oesophagus F(3, 31) = 61.15,

P<0.001; arms F(3, 31) = 32.53, P<0.001), with an average pHe for both regions of pHe 6.6 in ambient, pH 7.8, and pH 7.4 respectively. The pHe in treatment 7.6 in the oesophagus and arms was significant lower by 0.6 and 0.48 pH units respectively when compared to the other three treatments Figure 3.14 & 3.15, Table 3.4).

Extracellular pH in P. huttoni gut varied between pHe 6.62 and pHe 6.86 across treatments (Figure 3.16 & 3.17, Table 3.5) with an average pHe of 6.67 (n= 9, SE= 0.03) under ambient conditions. pHe was significantly lower in the gut in treatment pH

7.4 compared with the ambient pH treatment (F(2, 28) = 562.19, P<0.001). The oesophagus and arms regions for all pH treatments measured a pHe of 6.63, with no significant differences among pH treatments (Figure 3.16 & 3. 17, Table 3.5).

Extracellular pH in A. placenta gut varied between pHe 6.63 and pHe 8.2 with an ambient average of pHe 7.46 (n = 9, SE = 0.15) . Treatments pH 7.8, pH 7.6, and pH 7.4 were all significantly lower than ambient by 0.6 pH units, measuring a pHe of 6.89

(F(3, 35) = 5.12, P<0.005). The oesophagus showed small variation among treatments with no visible trend, pHe was 6.9 ± 0.1 for all treatments. Rod pHe was also very robust and measured the same pHe (6.9 ± 0.1) for all treatments. There was no significant effect of pH on pHe in the oesophagus or arms region (Figure 3.18 & 3.19, Table 3.6).

Extracellular pH in S. neumayeri blastocoel varied between pHe 6.1 and pHe 6.39 across treatments. The mean pHe for all treatments was pH 6.3 ± 0.02 (Figure 3.20 &

3.21, Table 3.7). There was no significant effect of external pH treatment on pHe (F(3,

39) = 0.89, P>0.45). The blastoderm pHe was lower than the blastocoel, measuring pHe of 5.9 ± 0.04 across all treatments. There was, however, no significant effect of pH treatment on pHe (F(3, 39) = 1.88, P = 0.15) (Figure 3.20 & 3.21, Table 3.7). S. neumayeri pluteus pHe showed a lower pHe with decreasing seawater pH across all larval regions. Gut pHe varied between 6.16 and 7.96 with a mean of 6.96 (n = 10, SE = 0.14) for the ambient treatment (Figure 3.22 & 3.23, Table 3.8). Mean pHe in the lowest pH treatment was significantly lower than in ambient pH seawater (F(3, 39) = 3.79, P<0.01) (Table 3.8). The oesophagus pHe varied between pHe 6.11 and pHe 6.53 across treatments with an ambient pHe average of 6.46 ± 0.03. Mean pHe in the lowest treatment was significantly lower than in the ambient pH treatment (F(3, 39) = 3.10, P < 0.05) . The rod pHe was similar to the oesophagus and varied between 6.11 and 6.57 with an ambient average of pHe 6.49 (F(3, 39) = 3.52, P < 0.05) (Figure 3.22 & 3.23, Table 3.8).

Extracellular pH in O. validus gastrula body cavity (BC) showed a decreasing trend as seawater pH decreased, pHe varied between 6.23 and 6.3 with an ambient average of 6.28 (n = 10 SE = 0.002) (Figure 3.24 & 3.25, Table 3.9). pHe in the lowest treatment was significantly lower than ambient by 0.4 pH units (F(3, 39) = 5.0, P < 0.005) (Table 3.9). In the gut, pHe was higher than in the BC in all treatments. pHe varied between 6.28 and 6.3 and pHe in treatment 7.2 was significantly lower than the other three treatments (F(3, 39) = 22.56, P < 0.001) (Figure 3.24 & 3.25, Table 3.9). O. validus bipinnaria pHe in the gut varied between pH 6.28 and pH 6.3 with an ambient average of pH 6.29 (n = 10 SE = 0.003) (Figure 3.26 & 3.27, Table 3.10). In the oral hood, mean pHe in treatment pH 7.2 was significantly lower than in pH 7.5 (F(3, 39) = 4.408, P < 0.01), but was not different from ambient pH or pH 7.7 (Table 3.10). In the gut pHe also varied between pH 6.28 and pH 6.3 with an ambient average of 6.29. Also, pHe in treatment 7.2 was significantly lower than in pH 7.7 (F(3, 39) = 11.78, P < 0.001). Conversely the oesophagus was reasonably consistent and pHe measured 6.29 (2.dp) across all treatments, thus pH did not have a significant effect on pHe in the oesophagus region (Figure 3.26 & 3.27, Table 3.10).

As with all species, the gut showed the highest variation in pHe among seawater pH treatments. For all species gut pHe of ambient treatment pH was significantly higher than pH in the three lower pH treatments (with the exception of O. validus bipinnaria).

Ambient pH 7.8 pH 7.6 pH 7.4

Oesophagus Gut Arm

Figure 3.14 Images of 12 day old Evechinus chloroticus larvae and resulting false colour pH map produced in MATLAB when raised under varying seawater pH. Scale bar represents100µm.

Table 3.4 One-way ANOVA for Evechinus chloroticus pluteus to identify significant differences in pHe among pH treatments. Body components tested are gut (a), gut wall (b), oesophagus (c) and arms (d). Data were log transformed and a Bartlett test confirmed homogeneity of variance. Heterogeneous variances were compared using Welch ANOVA testing for equal means, allowing for unequal variances. a) Gut Source DF SS MS F-ratio p pH treatment 3 0.036 0.012 34.56 <.0001 Error 28 0.009 0.0003 C. Total 31 0.047 Bartlett test's (F = 7.12, P = < 0.001)

Welch (F(3, 31)=139.77, P < 0.0001)

71 b) Gut wall Source DF SS MS F-ratio p pH treatment 3 0.0077 0.0026 18.86 <.0001 Error 28 0.0038 0.0001 C. Total 31 0.011 Bartlett test (F = 2.82, P = 0.037)

Welch (F(3, 31) = 13.06, P < 0.001) c) Oesophagus Source DF SS MS F-ratio p pH treatment 3 0.012 0.0041 61.15 <.0001 Error 28 0.002 0.00006 C. Total 31 0.014 Bartlett test (F = 2.867 p= 0.0351)

Welch (F(3,31) 52.35 P < 0.0001) d) Arms Source DF SS MS F-ratio p pH treatment 3 0.0061 0.002 32.53 <.0001 Error 28 0.0018 0.000063 C. Total 31 0.0079 Bartlett test (F = 7.72, P < 0.0001)

Welch (F(3,31) = 12.34 p=0.0003)

a) b) Gut Gut wall a b 7.8 7.8 7.3 c 7.3 a c ab 6.8 6.8 b pHe ± SE ± pHe

pHe ± SE ± pHe c 6.3 6.3 5.8 Ambient 7.8 7.6 7.4 5.8 Ambient 7.8 7.6 7.4 Seawater pH Seawater pH

c) d) Oesophagus Arms

7.8 7.8

7.3 7.3 a a a a a a 6.8 6.8 b pHe ±SE pHe pHe ± SE ± pHe 6.3 b 6.3

5.8 5.8 Ambient 7.8 7.6 7.4 Ambient 7.8 7.6 7.4 Seawater pH Seawater pH

Figure 3.15 Mean extracellular pH in Evechinus chloroticus under ambient pH and three reduced seawater pH levels for the gut (a), gut wall (b), oesophagus (c) and post oral arms (d). Error bars are ± standard error. Significant differences among pH treatments are indicated by columns not sharing the same letter. n=8.

Ambient pH 7.8 pH 7.6 pH 7.4

Gut Oesophagus Arm

n/a

Figure 3.16 Images of 12 day old Pseudechinus huttoni larvae and resulting false colour pH map produced in MATLAB when raised under ambient pH and three reduced seawater pH levels. Scale bar represents 100µm.

Table 3.5 One-way ANOVA for Pseudechinus huttoni pluteus to identify significant differences in pHe among pH treatments. Body components tested are gut (a), oesophagus (b) and arms (c). Data were log transformed and a Bartlett test confirmed homogeneity of variance. Heterogeneous variances were compared using Welch ANOVA testing for equal means, allowing for unequal variances.

a) Gut Source DF SS MS F-ratio p pH treatment 2 0.013 0.0064 562.19 <0.001 Error 26 0.0003 0.000011 C. Total 28 0.013 Bartlett test's N/A (all means were the same)

b) Oesophagus Source DF SS MS F-ratio p pH treatment 2 3.23e-7 1.61e-7 0.86 0.43 Error 26 4.87e-6 1.87e-7 C. Total 28 5.19e-6 Bartlett test (F = 1.655, P > 0.89)

c) Arms Source DF SS MS F-ratio p pH treatment 2 1.26e-6 6.32e-7 2.78 0.08 Error 26 5.90e-6 2.33e-7 C. Total 28 7.17e-6 Bartlett test (F = 2.99, P > 0.05)

Welch (F(2, 28) = 3.44, P > 0.057)

b) a) Gut Oesophagus a 6.7 6.7 a b 6.6 6.6 6.5 6.5 6.4 6.4 ±SE pHe pHe ± SE ± pHe 6.3 6.3

6.2 6.2 Ambient 7.8 7.4 Ambient 7.8 7.4 Seawater pH Seawater pH

c) Arms 6.7

6.6

6.5

6.4 pHe ± SE ± pHe 6.3

6.2 Ambient 7.8 7.4 Seawater pH

Figure 3.17 Mean extracellular pH in Pseudechinus huttoni under ambient pH and three reduced seawater pH levels for the gut (a), oesophagus (b) and arms (c). Error bars are ± standard error. Significant differences among pH treatments are indicated by columns not sharing the same letter. n=10.

Ambient pH 7.8 pH 7.6 pH 7.4

Oesophagus

Arm Stomach

Figure 3.18 Images of four day old Arachnoides placenta larvae and resulting false colour pH map produced in MATLAB when raised under ambient pH and three lowered seawater pH levels. Scale bar represents100µm.

Table 3.6 One-way ANOVA for Arachnoides placenta pluteus to identify significant differences in pHe among pH treatments. Body components tested were gut (a), oesophagus (b) and arms (c). Data were log transformed and a Bartlett test confirmed homogeneity of variance. Heterogeneous variances were compared using Welch ANOVA testing for equal means, allowing for unequal variances. a) Gut Source DF SS MS F-ratio p pH treatment 3 0.008 0.0026 5.12 < 0.01 Error 32 0.016 0.0005 C. Total 35 0.024 Bartlett test's (F = 0.829 P > 0.4774)

77 b) Oesophagus Source DF SS MS F-ratio p pH treatment 3 0.0009 0.0003 1.78 0.17 Error 32 0.0057 0.0002 C. Total 35 0.0066 Bartlett test's (F = 1.09 P > 0.354) c) Arms Source DF SS MS F-ratio p pH treatment 3 0.0010 0.0003 2.02 0.13 Error 32 0.0054 0.0002 C. Total 35 0.0064 Bartlett test's (F= 0.38 P > 0.7645)

a) b) Gut Oesophagus 7.7 a 7.7

7.5 7.5 b 7.3 b b 7.3 pHe ±SE pHe pHe ±SE pHe 7.1 7.1

6.9 6.9

6.7 6.7 Ambient 7.8 7.6 7.4 Ambient 7.8 7.6 7.4 Seawater pH Seawater pH

c) Arms

7.7 7.5 7.3

pHe ± SE ± pHe 7.1 6.9 6.7 Ambient 7.8 7.6 7.4 Seawater pH

Figure 3.19 Mean extracellular pH in Arachnoides placenta under ambient pH and three reduced seawater pH levels for gut (a), oesophagus (b) and arms (c). Error bars are ± standard error. Significant differences among pH treatments are indicated by columns not sharing the same letter. n=9.

Ambient pH 7.7 pH 7.5 pH 7.2

Primary mesenchyme cells Blastoderm Blastocoel

Figure 3.20 Images of seven day old Sterechinus neumayeri embryos and resulting false colour pH map produced in MATLAB when raised under varying seawater pH. Components tested were the blastocoel (a) and blastoderm (b). Scale bar represents 50µm.

Table 3.7 One-way ANOVA for Sterechinus neumayeri blastula to identify significant

differences in pHe among pH treatments. Body components tested were the blastocoel (a) and blastoderm (b). Data were log transformed and a Bartlett test confirmed homogeneity of variance. Heterogeneous variances were compared using Welch ANOVA testing for equal means, allowing for unequal variances.

a) Blastocoel Source DF SS MS F-ratio p pH treatment 3 0.0005 0.00017 1.88 0.15 Error 36 0.0034 9.4e-5 C. Total 39 0.0039 Bartlett test (F = 2.14, P > 0.13) b) Blastoderm

Source DF SS MS F-ratio p pH treatment 3 4.96e-5 0.00002 0.89 0.45 Error 36 0.0006 1.8e-5 C. Total 39 0.0007 Bartlett test (F = 0.99, P > 0.41)

a) Blastocoel b) Blastoderm

6.30 6.30 6.20 6.20 6.10 6.10 pHe ±SE pHe 6.00 ±SE pHe 6.00 5.90 5.90 5.80 5.80 Ambient 7.7 7.5 7.2 Ambient 7.7 7.5 7.2 Seawater pH Seawater pH

Figure 3.21 Mean extracellular pH in Sterechinus neumayeri blastula under ambient pH and three reduced seawater pH levels for the blastocoel (a) and blastoderm (b). Error bars are ± standard error. Significant differences among pH treatments are indicated by columns not sharing the same letter. n=10.

Ambient pH 7.7 pH 7.5 pH 7.2

Oesophagus Sterechinus neumayeri (4 weeks) Arm Gut

Figure 3.22 Images of 4 week old Sterechinus neumayeri larvae and resulting false colour pH map produced in MATLAB when raised under varying seawater pH. Scale bar represents 50µm.

Table 3.8 One-way ANOVA for Sterechinus neumayeri pluteus to identify significant differences in pHe among pH treatments. Body components tested were the gut (a), oesophagus (b) and arms (c). Data were log transformed and a Bartlett test confirmed homogeneity of variance. Heterogeneous variances were compared using Welch ANOVA testing for equal means, allowing for unequal variances. a) Gut Source DF SS MS F-ratio p pH treatment 3 0.0046 0.0015 3.79 0.018 Error 36 0.014 0.0004 C. Total 39 0.019 Bartlett test (F = 1.45, P > 0.226) b) Oesophagus Source DF SS MS F-ratio p pH treatment 3 0.00044 0.00015 3.11 0.038 Error 36 0.0017 4.75e-5 C. Total 39 0.0022 Bartlett test (F = 0.94, P > 0.42) c) Arms Source DF SS MS F-ratio p pH treatment 3 0.00044 0.00015 3.52 0.025 Error 36 0.0015 0.000042 C. Total 39 0.0019 Bartlett test (F = 2.23, P > 0.08)

a) Gut b) Oesophagus a a ab 7.10 7.10 ab ab ab b 6.90 b 6.90 6.70 6.70 pHe ±SE pHe pHe ± SE ± pHe 6.50 6.50

6.30 6.30 Ambient 7.7 7.5 7.2 Ambient 7.7 7.5 7.2 Seawater pH Seawater pH

c) Arms

7.10

6.90

6.70

pHe ±SE pHe a ab ab 6.50 b 6.30 Ambient 7.7 7.5 7.2 Seawater pH

Figure 3.23 Mean extracellular pH in Sterechinus neumayeri under ambient pH and three reduced seawater pH levels for the gut (a), oesophagus (b) and arms (c). Error bars are ± standard error. Significant differences among pH treatments are indicated by columns not sharing the same letter. n=10.

Ambient pH pH 7.7 pH 7.5 pH 7.2

Odontaster validus (15 days old) Oral hood Gut

Figure 3.24 Images of 15 day old Odontaster validus larvae and resulting false colour pH map produced in MATLAB when raised under varying seawater pH. Scale bar represents 100µm.

Table 3.9 One-way ANOVA for Odontaster validus gastrula to identify significant differences in pHe among pH treatments. Body components tested were the oral hood (a) and gut (b). Data were log transformed and a Bartlett test confirmed homogeneity of variance.

Heterogeneous variances were compared using Welch ANOVA testing for equal means, allowing for unequal variances. a) Oral hood Source DF SS MS F-ratio p pH treatment 3 0.000047 1.64e-5 22.56 <0.001 Error 36 2.52e-5 6.99e-7 C. Total 39 7.23e-5 Bartlett test (F= 1.92, P > 0.12) b) Gut Source DF SS MS F-ratio p pH treatment 3 2.73e-6 9.09e-7 5.01 <0.01 Error 36 6.56e-6 1.81e-7 C. Total 39 9.28e-6 Bartlett test (F = 0.53, P > 0.67)

a) Oral hood b) Gut a 6.30 6.30 a a 6.29 a 6.29 b a 6.28 a 6.28 6.27 6.27 6.26 6.26 pHe ± SE ± pHe 6.25 b SE ± pHe 6.25 6.24 6.24 6.23 6.23 Ambient 7.77.87.7 7.57.67.5 7.27.2 Ambient 7.77.8 7.57.6 7.2 Seawater pH Seawater pH

Figure 3.25 Mean extracellular pH in Odontaster validus gastrula under ambient and reduced seawater pH levels for the oral hood (a) and gut (b). Error bars are ± standard error. Significant differences among pH treatments are indicated by columns not sharing the same letter. n=10.

Ambient pH pH 7.7 pH 7.5 pH 7.2

Oesophagu Odontaster validus bipinnaria (43 days) s Gut Oral hood

Figure 3.26 Images of 43-day old Odontaster validus larvae and resulting false colour pH map produced in MATLAB when raised under varying seawater pH. Scale bar represents 100µm.

Table 3.10 One-way ANOVA for 43 day old Odontaster validus bipinnaria larvae to identify significant differences in pHe among pH treatments. Body components tested were the gut (a), oral hood (b) and oesophagus (c). Data were log transformed and a Bartlett test confirmed homogeneity of variance. Heterogeneous variances were compared using Welch ANOVA testing for equal means, allowing for unequal variances , a) Gut Source DF SS MS F-ratio p pH treatment 3 2.98e-6 9.93e-7 4.41 <0.01 Error 36 8.11e-6 2.25e-7 C. Total 39 1.11e-5 Bartlett test (F = 3.26, P < 0.0207)

Welch (F(3, 39) = 9.617, P< 0.0004) b) Oral hood Source DF SS MS F-ratio p pH treatment 3 7.44e-6 2.48e-6 11.78 <0.001 Error 36 7.58e-6 2.10e-7 C. Total 39 1.50e-5

Bartlett test (F = 3.68, P <0.0116)

Welch (F(3, 39) = 21.41, P < 0.0001) c) Oesophagus Source DF SS MS F-ratio p pH treatment 3 3.76e-8 1.25e-8 0.23 0.88 Error 36 1.99e-6 5.55e-8 C. Total 39 2.03e-6 Bartlett test (F = 0.381 P < 0.767)

a) b) Gut Oral hood 6.30 a a ab 6.30 ab ab b 6.29 bc c 6.29 6.28 6.28

6.27 6.27 pHe ± SE ± pHe pHe ± SE ± pHe 6.26 6.26

6.25 6.25 Ambient 7.7 7.5 7.2 Ambient 7.7 7.5 7.2 Seawater pH Seawater pH

c) Oesophagus 6.30

6.29

6.28

6.27 pHe ± SE ± pHe 6.26

6.25 Ambient 7.7 7.5 7.2 Seawater pH

Figure 3.27 Mean extracellular pH in 43 day old Odontaster validus bipinnaria larvae under ambient and reduced seawater pH levels for the gut (a), oral hood (b) and oesophagus (c). Error bars are ± standard error. Significant differences among pH are indicated by columns not sharing the same letter. n=10.

3.6 Discussion

The primary objective of this study was to examine the effects of near future ocean acidification scenarios (year 2100, 2100-2300 and >2300) (Table 1.1 chapter 1) on morphological and extracellular pH (pHe) changes during the development of echinoderm larval stages across a range of latitudes. In order to identify any latitudinal differences, five species were selected; the tropical sand dollar (Arachnoides placenta) two temperate sea urchins (Evechinus chloroticus and Pseudechinus huttoni), one Antarctic asteroid (Odontaster validus) and one Antarctic sea urchin (Sterechinus neumayeri). Extracellular pH was measured using the fluorescent indicator HPTS, which allowed visualisation of extracellular pH in larvae which had been raised under various seawater pH.

These findings provide insight into how species from different latitudes may respond developmentally and physiologically to ocean acidification and help us visualise their ability to regulate internal pHe at different developmental stages and in different body regions of the larvae.

3.6.1 Adjustment of seawater pH and ambient pH

The expected increase in CO2 emissions may lead to a reduction in ocean pH by up to 0.77 units by the year 2300 (Caldeira and Wickett, 2003, Raven et al., 2005). To carry out experiments on the effects of ocean acidification on echinoderms we lowered seawater pH according to the projected ρCO2(aq) levels (Bernstein et al., 2008). The carbonate chemistry of the seawater was also described based on measured pH, salinity, temperature, and alkalinity for each site. There were latitudinal differences in the seawater ρCO2; unexpectedly Townsville had the highest ambient carbonate concentration of 529 µatm (25°C seawater - pH 8.08), although this may reflect the regional affects of where the Aims marine laboratory is located. McMurdo had the second highest carbonate concentration of 407 - 413 µatm (-1.9°C seawater - pH 8). Finally, Otago Harbour measured 349 - 383 µatm (9-15°C seawater - pH 8.16). These results were similar to recent studies, with measurements for Townsville of 526 µatm (Gonzalez-Bernat et al., 2013b), McMurdo of 410 µatm (Yu et al., 2013) and the

Otago harbour results only slightly higher than Gonzalez-Bernat (2013) study of 337 µatm.

Projected ρCO2 values in treated seawater reached as high as 3710 µatm; this condition acted as an extreme treatment for the polar experiments (resulting in pH 7.2). Undersaturation of calcite and aragonite in the polar experiments occurred from a reduced seawater pH of 7.7 and below. In the temperate experiments an undersaturation of calcite and aragonite was seen in the lowest pH treatment (pH 7.4) and undersaturation with respect to aragonite in pH 7.6. There was no carbonate undersaturation for the tropical experiment at any reduced seawater pH. All lowered seawater pH treatments used in this study were within the range of previous studies (Dupont et al., 2010, Clark et al., 2009, Ericson, 2010, Gonzalez-Bernat et al., 2013a).

Due to the cooler water temperatures, undersaturation of calcite and aragonite will occur earlier in polar waters. In the sub-Antarctic and polar regions, aragonite and calcite under-saturation is predicted to occur when ρCO2 reaches ~600 and ~900 µatm (Kleypas et al., 2006, McNeil and Matear, 2008) perhaps as early as 2050 in the Antarctic winter (McNeil and Matear, 2008). However, lower latitude (tropical, subtropical) regions are still expected to remain saturated with respect to aragonite and calcite by the end of the century (Fabry et al., 2008).

3.6.2 Extracellular pH measurements in echinoderms

Acid-base regulation occurs at extracellular, cellular and sub-cellular levels. It has been widely accepted that pH in different body compartments plays a key role in the maintenance of physiological function and limitations under environmental stress (Riebesell et al., 2010). For ocean acidification research, acid-base parameters should therefore be determined in relevant compartments of whole organisms acclimated to

CO2 levels according to ocean acidification scenarios (Pörtner et al., 2010). For this study the pluteus larvae was divided into three main compartments; gut, arms, and oesophagus (in the E. chloroticus study the gut wall also was defined).

Pluteus pHe in all larval regions was lower than the surrounding seawater ranging from a pHe of 5.85 to 8, depending on the species and body compartment. Our results show that pHe in P. huttoni and A. placenta are more tolerant to a decrease in seawater pH,

88 whereas E. chloroticus is the least tolerant, especially in the gut and gut wall which showed the greatest amount of pHe change. Both polar species showed a decreasing pHe across all body compartments as seawater pH decreased, but only in the lowest treatment was it significantly different to ambient. These results are consistent with many studies which indicate that the tipping point for many echinoid larval stages occurs at the lower or more extreme pH treatments ≤ pH 7.4 (Clark, 2009, Ericson, 2010, Gonzalez Bernat, 2011, Kurihara et al., 2004).

Previous measurements of pHe

Recent studies have revealed that the pH in the stomach of the sea urchin, Stronylocentrotus droebachiensis larvae is alkaline (pH 9.5) and higher than any other body components (Stumpp et al., 2013). Alkaline conditions favour the breakdown of plant materials; algal and plant chloroplasts are more efficiently digested at a higher pH than at acidic or neutral pH (Schwenzfeier et al., 2011). Gastric pH decreased by up to 0.5 pH units when larvae were exposed to pH 7.4, indicating alkalinity could not be maintained resulting in a decreased efficiency to digest food. This decrease in efficiency then triggered an increase in feeding to compensate for energy loss (Stumpp et al., 2013). Thus, additional costs for acid-base regulation to maintain alkaline digestive conditions or other energy consuming compensatory digestive functions (for example, increased nutrient absorption) could significantly affect the organism's energy budget (Stumpp et al., 2013). The results in the current study were consistent with Stumpp et al., (2013), with the gut measuring a higher pHe than any other body components, although all my gut pH measurements were lower than pH 9 (pH 6.3 -7.9). When exposed to lowered seawater pH, pHe also decreased, also similar to Stumpp et al., (2013).

Earlier studies by Stumpp et al. (2012), found via measuring pHe with a micro- electrode, that pHe of Strongylocentrotus droebachiensis larvae conformed to the pH of the surrounding seawater (Figure 3.28 a). Although the micro-electrode is an effective method of measuring internal pH, it does however have a few pitfalls, such as imprecise calibration and differences in the ionic composition of that of buffers and body fluids. Measuring small animals, such as larvae, decreases the tip diameter of the sensor which can decrease overall sensitivity and accuracy (Riebesell et al., 2010).

In adult echinoderms (and some pluteus larvae), a number of authors have noted that

the CO2 content of the coelomic fluid is higher than that of the surrounding sea water, due to the tube feet being the primary medium for gas exchange (Table 3.11) (Miles et al., 2007, Spicer, 1995, Spicer et al., 1988). For example, the coelomic fluid pH of adult Paracentrotus lividus was observed to be lower than that of the surrounding seawater and decrease with decreasing seawater pH (Figure 3.28 b) (Catarino et al.,

2012). Elevated ρCO2 levels in echinoderms is considered to be responsible for reductions in pHe of 0.5 to 1.5 pH units compared with seawater, for example, the pHe of coelomic fluid from adult Psammechinus miliaris and Echinus esculentus was lower

than the surrounding ambient seawater (measuring pHcf 7.05 to 7.17) (Spicer et al., 1988). Further studies revealed that under acidified seawater conditions Psammechinus miliaris coelomic fluid pH also decreased with a decrease in seawater pH (Spicer et al. 1995) (Figure 3.28 c) Other marine invertebrates have provided similar results; hypercapnia caused a respiratory acidosis of extracellular fluids in the mussel Mytilus galloprovincialis resulting in a permanent reduction of pHe from 7.55 ± 0.02 to 7.36 ± 0.05 (Michaelidis et al., 2005).

In comparison with the results of this study with previous measurements of extracellular pH, the overall pHe values I measured were slightly lower than that of previous studies using adult echinoderms and also in Strongylocentrotus droebachiensis pluteus larvae (Table 3.11 and 3.12).

Table 3.11 Measurements of extracellular pH from four adult sea urchins, one pluteus sea urchins and one adult sea star, when reared under ambient seawater

conditions.Species Stage pHe Technique Source Psammechinus miliari Adult 7.4 (0.05) Micro electrode Miles et al., s 2007 Strongylocentrotus Pluteus 7.76 (0.06) Micro electrode Stumpp et al., droebachiensis 2012b Psammechinus miliaris Adult 7.10 (0.04) Micro electrode Spicer et al., 1988 Echinus esculentus. Adult 7.06 (0.03) Micro electrode Spicer et al. 1988 Strongylocentrotus Adult 7.55 (0.02) Micro electrode Dupont and droebachiensis Thorndyke (2012) Leptasterias polaris Adult 7.64 (0.01) Micro electrode Dupont and (seastar) Thorndyke (2012)

a) b)

pH (CF) pH pH (NBS)pH

Seawater pH (NBS) Time (hours) c)

0 1 2 3 4 5 Time (hr)

Figure 3.28 a) Relationship between seawater and the primary body cavity pH (NBS scale) of Strongylocentrotus droebachiensis pluteus during acute and chronic environmental hypercapnia. Bars represent ±SD; n = 4–5. (Stumpp et al., 2012 p.18193) b) Coelomic fluid pH (pH CF) of Paracentrotus lividus according to the aquarium se awater pH (pHSW) (Catarino et al., 2012 p 2348). c) Changes in pH in the coelomic fluid after exposure to hypoxia in Psammechinus miliaris, data are means + SD (n = 6) ( = coelornic fluid; = surrounding water) (Spicer et al., 1995 p 73).

Table 3.12 pHe values of the pluteus/bipinnaria regions (gut, oesophagus and arms/oral hood) of: a) A. placenta (n = 10), b) E. chloroticus (n=8), c) P. huttoni (n=10), d) Sterechinus neumayeri (n=10) and e) Odontaster validus when exposed to lowered seawater pH. Number in brackets indicated standard error.

Ambient(pH 8.1) pH 7.8 pH 7.6 pH 7.4 a) Arachnoides placenta Gut 7.46 (0.15) 6.86 (0.13) 6.87 (0.08) 6.88 (0.09) Oesophagus 6.84 (0.07) 7.02 (0.09) 6.99 (0.05) 6.85 (0.07) Arms 6.79 (0.06) 6.82 (0.08) 6.99 (0.07) 6.79 (0.06) b) Evechinus chloroticus Gut 7.91 (0.05) 7.42 (0.17) 6.5 (0.1) 6.72 (0.02) Oesophagus 6.59 (0.06) 6.64 (0.04) 5.97 (0.04) 6.64 (0.07) Arms 6.65 (0.02) 6.62 (0.03) 6.17 (0.013) 6.64 (0.067) c) Pseudechinus huttoni Gut 6.67 (0.03) 6.66 (0.002) n/a 6.64 (0.003) Oesophagus 6.64 (0.007) 6.64 (0.002) n/a 6.64 (0.002) Arms 6.63 (0.008) 6.63 (0.001) n/a 6.64 (0.001) d) Sterechinus neumayeri pH8.0 pH 7.7 pH 7.5 pH 7.2 Gut 6.96 (0.15) 6.65 (0.09) 6.60 (0.07) 6.52 (0.07) Oesophagus 6.46 (0.02) 6.43 (0.04) 6.4 (0.03) 6.33 (0.04) Arms 6.50 (0.02) 6.43 (0.04) 6.44 (0.02) 6.36 (0.04) e) Odontaster validus bipinnaria Gut 6.29 (0.003) 6.29 (0.001) 6.30 (0.001) 6.29 (0.001) Oesophagus 6.30 (0.001) 6.30 (0.001) 6.30 (0.001) 6.30 (0.002) Oral hood 6.30 (0.001) 6.30 (0.001) 6.30 (0.001) 6.30 (0.001)

3.6.3 Morphological observations with lowered pH

Data from this study confirms previous observations that lowering seawater pH alters the development and morphology of echinoderm larvae. Both early cellular development and pluteus/bipinnaria larvae growth showed either a delay in development or abnormal cell growth with a lowering of seawater pH. Extreme reductions in pH of 7.4 (tropical and temperate) and pH 7.2 (polar species) resulted in the smallest and most abnormal embryos and larvae.

Other studies have found that under increased seawater ρCO2(aq) concentrations, echinoderm larvae from tropical, temperate and polar regions are affected to varying degrees during different life stages. The main effects of lowered pH have been reduced larval size, altered body morphology, lowered calcification and lowered survival rates as development continues (Brennand et al., 2010, Clark et al., 2009, Ericson et al., 2010, Gonzalez-Bernat et al., 2013b, Kurihara et al., 2004, Pecorino, 2012) (Table 3.13). Larval abnormalities were observed under reduced oceanic pH levels for two species of sea urchins Hemicentrotus pulcherrimus and Echinometra mathaei (Kurihara and Shirayama, 2004). Furthermore, delays in the development rate were observed in Hemicentrotus pulcherrimus where fertilised eggs under ambient pH had reached the 8- cell stage while some remained at the 1-cell stage under lowered seawater pH (Kurihara and Shirayama, 2004). Developmental abnormalities have also been observed in other marine invertebrate groups including a decrease in shell length in the green lipped mussel Perna canaliculus when reared under lowered pH (Ericson, 2010). Also, when embryos of the oyster Crassostrea gigas were exposed to 2268 µatm ρCO2, after 24 hours more than 80% of larvae grown in high-CO2 seawater displayed malformed shells or remained unmineralized (Kurihara et al., 2007).

In this study, A. placenta showed the greatest number of abnormal embryos in the early developmental stages, additionally for the pluteus stage all body components were significantly reduced in length with decreasing seawater pH, consistent with Gonzalez- Bernat (2013b) results. My results for O. validus are also consistent with Gonzalez- Bernat (2011a) who also found that larvae raised under lowered pH were smaller and less developed. In contrast to studies by Byrne et al., (2013b) who found a significant reduction of normal pluteus development at pH 7.8 and 7.6, the morphometrics of S. neumayeri was the least affected in this study, however increased abnormality meant

93 the larval width increased with a lowering of pH, this was mainly due to the development of the arms which occurred in an abnormal direction under lowered seawater pH. Previous studies have found that S. neumayeri post-oral rod growth to reduce by 19%, an increase in left-right asymmetry and altered body allometry due to a decreased in pH which disrupts the natural development under lowered pH (7.6 and 7.8) (Byrne et al., 2013b). Our results found a 12% and 11% reduction in the post oral rods 1 and 2 respectively when raised in pH 7.4, but no evidence of left-right asymmetry in any species.

Polar larval development is extremely slow; Sterechinus neumayeri larvae take five days to hatch, ten days to reach gastrula, early pluteus is reached at 21 days and metamorphosis into a juvenile does not occur till 115 days (Bosch et al., 1987). The sea star, Odontaster validus has an even longer larval developmental process, it takes seven days to reach gastrula and up to 55 days to reach the bipinnaria stage where they then remain pelagic near the seabed for up to six months before they undergo metamorphosis into a juvenile starfish (Pearse, 1969). Due to this long time to complete each developmental stage, morphological delays in development may appear less distinguishable, but be more influential during these early stages.

As mentioned, sea urchin larvae exposed to decreased seawater pH have a decreased digestive efficiency (Stumpp et al., 2013), which in turn may decrease energy input causing a delay in growth. Hypercapnia can also induce metabolic depression, an adaptive strategy for survival under unfavourable conditions (Michaelidis et al., 2005). This lowered metabolism has been seen in the mussel Mytilus galloprovincialis which resulted in delayed growth, possibly due to the available energy being allocated to the maintenance of acid base regulation instead of growth (Michaelidis et al., 2005). Although delays in development and abnormalities were noted in this study not all species showed a significantly lower pHe in the lowest pH treatment. For example, in P. huttoni and A. placenta, larval width, length and both post-oral rod lengths were significantly reduced under pH 7.4, yet pHe was only significantly lower than ambient in the gut. The regulation of pHe and pHi may affect the cost of growth, in the lowest pH treatment of some species the regulation of internal pH takes priority so energy allocation is drawn away from growth to support the needs of acid base regulation (Michaelidis et al., 2005).

In contrast to the planktotrophic larvae in this study there has been evidence that lecithotrophic echinoderm larvae and juveniles are positively impacted by ocean acidification, previously observed in the sea star Crossaster papposus which grew faster under lowered pH seawater, with no visible effects on survival or skeletogenesis (Dupont et al., 2010a). This finding suggests lecithotrophic species may be better adapted to deal with the threat of ocean acidification compared with planktotrophic ones, for example, Heliocidaris erythrogramma (a lecithotropic sea urchin) showed no difference in normal development under lowered seawater pH (Byrne et al., 2009). This higher tolerance could possibly be because they have more energy available from the yolk. This may have potentially important consequences at the ecosystem level (Dupont et al., 2010a).

Table 3.13 Examples of the effects of reduced seawater pH on the different early developmental stages and survival of a range of marine calcifiers from tropical, temperate and polar regions.

Taxon Species Climate Effect of OA Source

Fertilisation Blastula/gastrula Pluteus Survival Echinoid Arachnoides placenta Tropical Decreased Delayed Reduced size Decreased (Gonzalez-Bernat et al., 2013b) Gastropod Haliotis coccoradiata Tropical Normal Delayed Reduced numbers Decreased (Byrne et al., 2010b)

Echinoid Echinometra mathaei Tropical Decreased Abnormal Reduced size/ - (Kurihara and abnormal Shirayama, 2004) Echinoid Hemicentrotus pulcherrimus Tropical Decreased Delayed/ reduced Reduced size Decreased (Kurihara and numbers (adult) Shirayama, 2004) (Shirayama and Thornton, 2005) Echinoid Tripneustes gratilla Tropical Normal - Reduced size Decreased (Clark et al ., 2009) (Byrne et al., 2010b)

Asteroid Patiriella regularis Temperate Normal Delayed/abnormal Reduced growth Decreased (Byrne et al., 2013a)

Echinoid Evechinus chloroticus Temperate - Delayed Reduced size and Decreased (Clark et al., 2009) calcification Echinoid Pseudechinus huttoni Temperate - Delayed Reduced size and Decreased (Clark et al., 2009) calcification Ophiuroid Ophiothrix fragilis Temperate Decreased Abnormal Delayed, Abnormal Decreased (Dupont 2008)

Bivalve Perna canaliculus Temperate - - Reduced size and - (Ericson 2010) calcification

Asteroid Odontaster validus Polar - Abnormal Reduced size Decreased (Gonzalez-Bernat et al., 2013a)

Nemertea Parborlasia corrugatus Polar Normal - Reduced size - (Ericson et al., 2010)

Echinoid Sterechinus neumayeri Polar Decreased Delayed Reduced size, normal Decreased (Clark et al., 2009) calcification (Ericson et al., 2010)

(Byrne et al., 2013b) Gastropod Limacina helicina Polar - - Reduced calcification Decreased (Comeau et al., 2009)

Crustacean Euphausia superba Polar Disrupted Abnormal - Decreased (Kawaguchi et al., 2011) development

3.6.4 Physiological responses

Acid/base regulation

Acid-base and metabolic regulation are inter-dependent processes, so that changes in pH can affect metabolic rate, the mode of catabolism, and energetic budgets, additionally influencing protein functioning and oxygen transport (Portner and Sartoris, 1999). To maintain high enzyme activities, digestive system pH is regulated by active ion transport processes through the net export or import of acid equivalents (Stumpp et al., 2013). One of the main acid-base regulation mechanisms in animals is associated with active ion transport (gills, renal or digestive tissues). Hypo-metabolic and osmoconfomers (such as sea urchins) are therefore less able to cope with ocean acidification because they have a less efficient ion regulatory machinery and buffers, both which could protect physiological fluids against hypercapnia (Catarino et al., 2012, Melzner et al., 2009, Portner, 2008).

Sea urchins are thus not good acid-base regulators and have a low concentration of ion regulating proteins in the coelomic fluids (Spicer et al., 1988). Under increased seawater acidity, hypercapnia in the coelomic fluid in the sea urchin Psammechinus miliaris was eventually shown to cause mortality (Miles et al., 2007). The Miles et al. (2007) study revealed that a reduction of seawater below pH 7.5 would be markedly detrimental to the acid-base balance of the sea urchin, regardless of its ability to tolerate fluctuations in pH in its environment. Miles et al. (2007) also tested P. miliaris in pH 6.16 which resulted in a pH drop in the coelomic fluid from pH 7.53 to pH 6.42, at this point, cells are more likely to rupture causing intra and extracellular leakage that would prevent any accurate measurements of coelomic fluid pH.

The maintenance of extracellular pH is crucial in protecting individuals against acidic disturbances that may alter development (Catarino et al., 2012). Skeletal growth is controlled by primary mesenchyme cells (PCM), a decrease in pHe directly affects the calcifying PCMs as it interferes with the pHi regulatory systems via decreasing proton gradients. The production of amorphous calcium carbonate (ACC) occurs within the PMCs and is directly linked to pHi regulation. pH homeostasis is therefore very important and the ability to maintain full functioning ion transporters to prevent inhibiting of any of these crucial acid-base regulating systems (Stumpp et al. 2012).

Acid base buffering

For vertebrates, acid-base buffering is achieved with the aid of bicarbonate, phosphates, plasma proteins and haemoglobin (Davenport, 1974), but in the coelom (or blood) of invertebrates a high buffering capacity is due almost entirely to the presence of proteins

(Spicer et al., 1988). And, as Spicer et al. (1988) found, the CO2 capacity of the coelomic fluid of two sea urchin species (Psammechinus miliaris and Echinus esculentus) was only marginally greater than that of seawater, which is consistent with their low protein concentration in the coelomic fluid. Stumpp et al. (2012) suggested that the extracellular space may act as a buffer so intracellular pH is maintained and processes (such as calcification) can proceed under more acidified environmental conditions, this is shown in the sea urchin Paracentrotus lividus which used its coelomic fluid as a buffer (Catarino et al., 2012). In the current study, pHe is consistently lower than the surrounding seawater pH, possibly due to the extracellular space acting as a buffer to protect intracellular processes as stated in the above study.

Additionally, some CO2 tolerant species, such as the sea urchin Psammechinus miliaris (Miles et al., 2007), have shown the ability to perform a pH compensatory reaction that protects their extracellular fluids from ocean acidification, important in avoiding metabolic depression (Melzner et al., 2009).

Sterechinus neumayeri showed a decreasing trend in pHe across all body regions when exposed to lowered seawater pH, although pHe in treatment pH 7.2 was only significantly lower than ambient. Acid-base regulation is energy dependent, which species are capable of modulating (e.g. decreasing energy for metabolism or growth to increase acid-base regulation). However, the rate and efficiency of acid-base regulation is influenced by the value of pHe that can be maintained by the ion exchange mechanisms (Reipschläger and Pörtner, 1996). Cold water species lack powerful ion exchange mechanisms (Reipschläger and Pörtner, 1996), this could explain why overall polar larvae had a lower pHe in all body compartments than tropical or temperate species. In addition to their extreme environment polar species may find it harder to maintain a consistent pHe, but only under extreme conditions of pH 7.2, as shown in my results.

Early cell development has also been shown to be sensitive to ocean acidification with many delays in cleavage rates. Protein synthesis is reduced due to a reduction in intracellular pH in zygotes incubated in lowered pH (Grainger et al., 1979), without extracellular compensation it is thought to lead to decreased growth and cleavage rates

(Kurihara, 2008). At concentrations greater than 5000 ppm CO2 there was a substantial decrease in the number of sea urchin zygotes capable of cleaving (Kurihara et al., 2004). Other studies have found mitotic abnormalities induced in sea urchin embryos at pH below 6.5 (Cipollaro et al., 1986, Pagano et al., 1985), a pH below the ability to most echinoderms to use extracellular compensation to aid intracellular activities (Stumpp et al., 2012b).

Impacts of compensation and ion regulation

Acid–base status and the capacity to regulate and compensate for acid–base disturbances are fundamentally important in setting sensitivity to ocean hypercapnia (Portner 2008). Acid-base regulation, however, not only means adjustment or maintenance of intra- and extracellular pH, but under certain conditions priority can be 3- - given to the regulation of the concentrations of bases (CO2 , HCO3 ) or acid (H2CO3) in body fluids (Pörtner et al., 2010). In this case, pHe then becomes a dependant variable (Riebesell et al., 2010), therefore making it possible that pHe levels decrease in order to compensate for pHi (Melzner et al., 2009). Miles et al., (2007) study on sea urchin - acid-base regulation recorded a series of increases in HCO3 in a zigzag pattern over a seven day period when they exposed adult urchins to lowered seawater pH, illustrating successive rises and falls in coelomic pH. These fluctuations could potentially occur when coelomic fluid pH gets too low (reaches a threshold), triggering an up-regulation -, of HCO3 which is used for acid base compensation.

Buffering of tissues has also been seen as another form of altering acid-base concentration and can be achieved through an increase in bicarbonate concentrations, which in calcified organisms, can be made through dissolution of calcite (such as the larval skeleton) (Clark et al., 2009). For example, mussels are able to support acid-base regulation by increasing haemolymph bicarbonate levels, derived from the dissolution of shell CaCO3 (Michaelidis et al., 2005). Consequently, this compensation makes

100 mussel shells weaker and more vulnerable to predation. The Clark et al. (2009) results showed pitted, eroded larval skeletons which may have also been due to acid-base regulation by increasing bicarbonate levels in the extracellular and intracellular space. In this study all larvae showed a drop in pHe as treatment pH was decreased, however, there may be a limited buffering capacity among species. To answer this question, we would need to investigate a way to measure pHi at the same time as pHe in order to see ion flow across the intra- and extracellular membranes.

In echinoderms, acid-base regulation is largely controlled through the important ion regulatory membrane-transporter Na+/K+-ATPase (Melzner et al., 2009). It has a high energy demand and may account for up to 80% of the total metabolic rate in some pluteus larvae (Leong and Manahan, 1997). pHi regulation under ambient conditions is achieved by ion exchange between intra- and extracellular compartments. It is likely that Na+/K+-ATPase activity is increased with increasing environmental acid–base stress through CO2 induced acidification (Stumpp et al., 2011b). This change in energy allocation may be a related to reduced growth in echinoderms, for example, Na+/K+- ATPase gene expression was up-regulated under hypercapnic conditions in purple sea urchin larvae (Strongylocentrotus purpuratus), there was also an increase in metabolic rate and developmental delays, indicating a higher energy demand under high CO2 seawater conditions (Stumpp et al., 2011b). In this study growth rates also decreased, possibly due to the increase in Na+/K+-ATPase activity in order to regulate pHi.

Another suggestion is that as pHe decreases, ion regulation through the Na/H+ + - + - exchanger is inhibited, and the more economic Na /HCO3 /H /Cl (which transports two acid-base equivalents per N+, instead of one) exchange becomes more active while reducing the activity of the energy demanding Na+K+-ATPase (Figure 3.29) (Pörtner and Bock, 2000). This has been supported in a study where larvae of the sea urchin Strongylocentrotus purpuratus significantly decreased their mRNA levels for the

Na+/K+- ATPase in response to high CO2 conditions (Todgham and Hofmann 2010).

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Decreases

Increases

pHi (Intracellular) pHe (Extracellular)

Figure 3.29 Ion regulation in invertebrate larvae during hypercapnia. When acidity increases, extracellular protons inhibit (-) the Na+/H+ exchanger. As a consequence, + + - - the Na /H /Cl /HCO3 exchanger becomes increasingly involved in pHi regulation while reducing in Na+/K+-ATPase activity (modified from Portner et al., 1998 p1806).

This transport exchange, in addition to reduced oxygen consumption, provides evidence that the ATP demand of acid-base regulation comprises a significant fraction of metabolic rate which falls during metabolic depression (Pörtner and Bock, 2000). The energy contribution to regulate acid-base homeostasis differs among species, with the active component of pH regulation thought to be much larger in eurythermal species (over 50% of pH change in the shrimp Crangon crangon), whereas in stenotherms (e.g., S. neumayeri) it equates to only 10% (Sartori & Portner 1997). Also unlike euyrthermal animals who mostly use active mechanisms of pHi adjustment, stenotherm's predominantly use passive mechanisms of ion transport which allows flexible adjustments of pH according to their metabolic requirements (Pörtner and Bock, 2000). Therefore, animals living at lower temperatures may be able to compensate for acid- base disturbances faster since cold adaptation could increase the capacity of pH regulatory mechanisms. Sartoris and Portner (1997) suggest that a larger active than passive component of acid-base regulation may be needed in order to colonize shallow coastal waters, allowing for a more flexible response with temperature changes.

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Whereas, temperature changes experienced by Antarctic stenotherms (e.g. seasonal) involves minor fluctuations (Sartoris and Portner 1997).

Low levels of available energy may compromise acid-base and ionic regulation. To save energy, animals may tolerate passive changes of pHi during periods of inactivity. A down regulation of ion exchange could lead to a reduction in pHi (Portner et al., 1998). In Sipunculus nudus, pHe is the key acid-base parameter instigating metabolic depression during acidosis, whereas metabolic depression is not induced by moderate (0.3 units) decreases in pHi (Reipschläger and Pörtner, 1996). In general, pHi is regulated at the expense of pHe. The reduction in pHe could counteract high acidifying rates of anaerobic metabolism and very probably support the reduction of ATP turnover observed during anaerobic conditions (Reipschläger and Pörtner, 1996)

In echinoderms, metabolic rates can increase by an order of magnitude prior to metamorphosis (Leong and Manahan, 1997), therefore the ability to regulate pH during periods of a high metabolic demand are very important for successful development. It is still unclear whether species that can tolerate ocean acidification will be able to maintain compensatory responses over time. Disruptions in pH may only be tolerated in the extracellular compartments or hemolymph (in crustaceans) only for short periods of time (hours), and hemolymph pH regulation is an important factor in maintaining oxygen supply (Whiteley, 2011). Thus, long-term compensation can negatively affect other physiological functions. In addition, it is unclear whether less tolerant species will be able to acclimatise, or even adapt, to the changes in seawater carbonate chemistry (Whiteley, 2011). Compensation from environmental disruption is more likely to involve strong iono- and osmoregulators, with well developed ion exchange mechanisms, such as, freshwater crustaceans (Procambarus clarkii ) which can survive considerable acidification of their freshwater habitats (Weber and Pirow, 2009) due to them possessing the mechanisms that enable them to compensate for acid–base disturbances; at least in the shorter term (Whiteley, 2011). Echinoderms, being poor iono- and osmoregulators have limited abilities to compensate for acid–base disturbances; this is compounded in slow-moving, relatively inactive species because they have low circulating protein levels and low buffering capacities. Species living in low-energy environments, such as polar and deep sea habitats, are particularly vulnerable, because they are metabolically limited with respect to environmental

103 change (Whiteley, 2011). The true impact of ocean acidification would depend on the metabolism of species and the ability of organisms to regulate acid-base balance in tissues surrounding important structures (Fabry et al., 2008).

Scope for growth & respiration

A delay in larval development would occur as energy is reallocated in an attempt to regulate intracellular pH (Michaelidis et al., 2005). This regulation would, in turn, increase energetic costs and lead to a shift in the larvae’s energy budget away from growth and focused on ion pumping to prevent acidosis. Studies by Stumpp et al.

(2011) revealed that larvae raised under high ρCO2 spent less energy on somatic growth (39% to 45%) when compared with control larvae which allocated between 78% and

80% of the available energy into growth. Elevated ρCO2 has led to an increased respiration rates 2.1-times the rate of respiration in ambient conditions (Stumpp et al., 2011b). Respiration rates are considered to be a general indicator for stress and essential in quantifying energy demands in larvae, they can also give information on the overall impact of environmental changes (Marsh and Manahan, 1999). Stress has been known to cause an up-regulation of metabolic genes and metabolism, leading to a higher energy demand (Stumpp et al., 2011a, Stumpp et al., 2012a). Concurrently occurring is the down-regulation of calcification related genes, which causes delayed growth characteristics (Stumpp et al., 2011a). A further study revealed it is the down- regulation of genes involved in both the sequestration and binding of Ca2+ for mineral deposition during spicule formation and skeletogenesis (Todgham and Hofmann, 2009). High energy amounts are spent on acid base regulation of PMC homeostasis additionally compromising the available energy for growth (Stumpp et al., 2011a).

The stunted and delayed growth shown in many previous studies (Table 3.13), including ours is a visual sign of the effects of lowered pH on morphology. Visual signs, however, may hide some of the sub-lethal and subtle physiological changes that are not reflected by morphological changes. Up until recently, pHe has not been easily quantified, but by using techniques such as the one in this study we can identify the effects that ocean acidification has on the extracellular physiology of larvae. For example, polar species have the additional disadvantage of living in a thermally stressful environment and low food concentrations making them metabolically limited,

104 therefore their ability to regulate extracellular pH may not be as good as temperate or tropical species which are not metabolically limited. Morphologically, the degree of delay in development may not be distinguished due to their prolonged larval growth during early stages of development but there may be significant differences in pHe. In this study there was not a distinguishable delay in development in S. neumayeri pluteus when reared under pH 7.2, however, pHe was significantly lower in the treatment pH 7.2 compared with ambient. On the other hand, in Odontaster validus, the opposite occured and there was a delay in development and lower pHe under lowered treatment pH.

3.6.5 Latitudinal differences

Temperature is one of the most important abiotic factors shaping marine ecosystems due to its impact on all biological processes (Portner et al., 1998). Development rates in Antarctic benthic invertebrates are between two and twenty times slower than temperate species (Peck, 2005). These extremely cold temperatures may also set a limit for the adjustment of pH regulation to changing temperatures in response to climate change (Portner et al., 1998). Many species that live at constant near-freezing temperatures (-1.9°C) of the Southern Ocean die of acute heat death at temperatures only a few degrees above their normal habitat temperatures (e.g. approximately +1.8°C) (Peck et al., 2008, Somero, 2010). Clark et al. (2009) suggested that calcification was more difficult in cold waters due to lower calcium carbonate saturation levels. These species live in an extreme environment, and many have evolved a specialised ability to survive and calcify at below 0°C temperatures, it has also been found that mitochondrial contents of muscles in cold water ectotherms are higher at low temperatures, which may increase metabolic rates to overcome environmental constraints (Lurman et al. 2012, Peck and Conway, 2000). Stenotherms, on the other hand, such as Sterechinus neumayeri larvae, minimize their metabolic rate and the aerobic capacity per milligram of mitochondrial protein, thereby minimizing oxygen demand (Portner 2002, Watson et al., 2014). Some species, however, may already be at the lower limit of calcite production and further restrictions due to ocean acidification may be unsustainable.

A recent review by Byrne et al. (2013) summarised 26 studies and concluded that there were differences in the calcification and growth of larval rod across latitudes, due to the

105 differences in calcite (Ωc) and aragonite (Ωa) saturation states (Byrne et al., 2013c).

Tropical species were seen to be more vulnerable to reduced Ωc, whereas polar species were more resilient (Figure 3.30). Polar species are likely to have adapted to the stable and naturally low carbonate ion concentrations characteristic of these waters over millions of years (McNeil and Matear, 2008). Clark et al. (2009) also found that the polar species, S. neumayeri, was less sensitive to lowered pHs when comparing larval skeleton degradation with temperate and tropical species.

Figure 3.30 Reduced growth of the postoral arm in echinoplutei in response to changes in calcite and aragonite saturation. The solid line is the regression line and the shaded area is 95% CI. Different colours represent geographical regions (Byrne ., 2013c p7).

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In this study, polar species were shown to have a consistently lower gut pHe under acidified conditions when compared to tropical or temperate species (Figure 3.31). When I compare the larval gut, oesophagus and arm pHe across latitudes, pHe in each larval region decreases as latitude increases (Figure 3.32). Additonally, at each latitude the gut pHe is greater than the oesohagus and arms which both have a similar pHe. Gut pH has been found to be more akaline than other larval areas as it increases the ability to digest and dissolve dietary protein (Stumpp et al., 2013). Areas such as the larval arms are associated with high calcification (Brennand et al., 2010), and therfore the low pHe experienced could support the theory that pH is lowered in the extracellular space to maximise calcification in intracellular space under an optimal pHi (Portner et al., 1998). Stumpp et al. (2012) suggests that maintained intracellular pH enables calcification to proceed despite decreased pHe.

8.5

8.0 Tropical 7.5 Temperate

7.0 Polar

Gut pHe Gut pHe 6.5

6.0

5.5

5.0 7.0 7.2 7.4 7.6 7.8 8.0 8.2 Experimental pH

Figure 3.31 Changes in gut pHe in response to altered seawater pH. Species were pooled into either tropical (A. placenta), temperate (E. chloroticus and P. huttoni) and polar (S. neumayeri and O. validus) latitudinal regions.

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8.0 7.8 7.6

7.4 Gut 7.2 Oesophagus 7.0 pHe (±SE) pHe 6.8 Arms 6.6

6.4 6.2 0 20 40 60 80 Equator s Latitude Figure 3.32 pHe versus latitiude for mean pHe in the gut, oesophagus and arms regions in the five study species. Error bars are ± 1SE.

Evidence from ocean acidification experiments have suggested that several taxa might react more sensitively to ocean acidification, while others remain surprisingly tolerant (Melzner et al., 2009). Based on this study’s pHe findings, the temperate P. huttoni showed the most tolerance to a lowering seawater pH as shown by its ability to maintain a consistent pHe (to ambient conditions) when treatment pH dropped, although the gut pH dropped in pH 7.4, the larval arms and oesophagus remained consistent. Arachnoides placenta also showed a high tolerance, although the gut pH was significantly lower in all treatments below ambient. These finding contradict other studies which have found tropical species to be more vulnerable to lowered seawater pH (Byrne et al., 2013c, Gonzalez-Bernat, 2011), although these studies did not investigate acid-base regulation but focussed on morphometric and growth data, in which case this study also concludes that the tropical A. placenta growth was affected to the greatest degree morphologically. The polar Sterechinus neumayeri blastula showed no significant drop in pH in the blastocoel or blastoderm under lowered pH, however, the four week old pluteus was resilient to pH change up until a tipping point at 0.8 pH units below ambient (pH 7.2), where a significantly lower pHe was recorded in the gut, oesophagus and arms of the pluteus, this species was also found to be resilient to lowered pH by Clark et al., (2009). Odontaster validus does not develop a skeleton until late development (post-bipinnaria), this may explain why all pHe values

108 were the same in each region, as it may not need to maintain a specific pHi until it is time to make a skeleton. It would be interesting to keep monitoring pHe until the development of a skeleton to see if this added pressure under acidified conditions caused any changes in pHe.

The faster metabolism and growth rate of the tropical species could explain the higher tolerance in some larval compartments and ability to regulate pHe under all acidified treatments. Species with slower metabolism, such as Antarctic species which already have a delayed cell development due the cold waters (Stanwell-Smith and Peck, 1998) showed an ability to regulate pHe in this study when minor pH changes are made (-0.5 pH units), but show signs of interrupted regulation when dropped to pH 7.2. They also showed an overall lowered pHe than lower latitude species.

Skeletal composition

Mineral composition of echinoderm skeletons differ with latitude, with higher latitudes having less Mg2+ incorporated into their skeletons (Byrne et al., 2013c). Previous studies have found via scanning microscopic examination that calcification in the arms of S. neumayeri larvae were not affected at pH 7.5, whereas the skeleton of temperate E. chloroticus larvae was pitted in pH 7.7 and below (Clark et al., 2009). In this study, Sterechinus neumayeri pHe was not affected until pH 7.2, yet E. chloroticus pHe was significantly lower in pH 7.6. The low Mg2+ content characteristic of polar species could aid in pHe homeostasis as a dissolving skeleton would put more stress on acid base regulation. The adult test of the tropical sea urchin Echinometra mathaei has up to 16% Mg2+ content compared to the polar Sterechinus neumayeri, having one of the lowest Mg2+ content of only 6% (McClintock et al., 2011). Skeletons with higher Mg2+ content are more vulnerable to reduced carbonate saturation. As saturation levels of both calcite and aragonite decrease as latitude increases, it makes sense for polar species to include less Mg2+ into their skeletons. Therefore, because of skeletal mineralogy tropical urchins are expected to be vulnerable to ocean acidification (Bischoff et al., 1987) although polar regions show a lower saturation state. This mineral composition could make S. neumayeri more resilient, as shown in this study, to dissolution during ocean acidification compared to tropical or temperate species.

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Chapter 4 General Discussion and Conclusions

4.1 Summary of Results

The primary focus of this MSc research was to examine the effects of ocean acidification on extracellular pH in the developing larvae of five important benthic echinoderms, the tropical sand dollar Arachnoides placenta, two temperate echinoderms Evechinus chloroticus, and Pseudechinus huttoni and the Antarctic echinoid Sterechinus neumayeri and asteroid Odontaster validus. A fluorescent probe method was refined that allowed me to look at extracellular pH in larvae that had developed under different seawater conditions (0.3 to 0.8 pH units below ambient). Inter-specific differences were examined to determine how responses varied with latitude.

The two working hypotheses for this study were (1) Under CO2 driven ocean acidification, echinoderm larvae will have a lower capacity to regulate their pHe, resulting in a decrease in pHe with decreasing seawater pH, and; (2) Responses will vary as a function of latitude and will be species-specific, with less resilience observed in the polar species because they live in naturally low carbonate saturated waters, and due to cold temperatures and low food availability they are metabolically limited. Hypothesis 1 was supported for some aspects of development, such as the sea urchin pluteus and sea star bipinnaria, however responses varied between developmental

110 stages and larval regions. Responses also varied among species, meaning that pHe response to lowered pH is species specific. Overall the lowest treatment (pH 7.2 or pH 7.4) had the greatest effect on larvae, causing delays in morphology, abnormal growth and significantly lowering pHe during larval development.

Hypothesis 2 was somewhat supported as the temperate sea urchin P. huttoni and tropical sand dollar A. placenta pHe were the most resilient to the lowered pH conditions, with only the gut regions showing a significant decrease when in acidified conditions. Overall polar species showed a lower pHe in every region and pHe significantly decreased in the lowest treatment of pH 7.2. Evechinus chloroticus unexpectedly showed the least tolerance to pH change, with a significant decrease in pHe in all larval regions under treatment pH 7.6.

The response of echinoderms to ocean acidification in the current study differs somewhat from that previously reported. Polar echinoderms, previously shown to be potentially more tolerant than tropical or temperate species (although most previous studies based their results on growth and morphology), were found to be less able to regulate pHe, however, this was only under the more extreme conditions. Environmental pressures (such as low temperature and low calcium carbonate saturation states) may indicate that the ability of acid-base regulation in decreased pH is restrained in species that already have environmental pressures associated with living at low latitudes.

Many studies have outlined the negative effects of ocean acidification on a number of species, their form, and function. These include effects on fertilisation (Gonzalez Bernat, 2011), development (Byrne et al., 2010b, Dupont et al., 2008, Kurihara, 2008), calcification (Clark et al., 2009, Ericson, 2010), metabolism (Melzner et al., 2009, Portner et al., 1998) and more recently the effects on intra- and extracellular pH (Stumpp et al., 2012a). The following sections discuss the ecological impacts of my findings, the inclusion of synergistic effects, effects on cold water species, further research considerations and conclusions.

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4.2 Ecological impacts

This study has shown that projected ocean acidification scenarios will have some effect on echinoderm larval growth and their ability to regulate pHe. These benthic invertebrates all play an important role in their environment. Decreased growth rates will have deleterious consequences on the survival of many species. During larval development, it is favourable to minimize development time. Planktonic mortality is high and increasing the time a larva spends in the water column increases the chance of predation and delays the opportunity to settle; late-settlers often experience the lowest survival and settlement is vital for recruitment (Dupont et al., 2010b, Dupont and Thorndyke, 2009). Additionally, many ecological processes are synchronized, for example, barnacle larvae are released in the plankton to coincide with the spring bloom. If the development of these nauplii larvae is delayed, synchrony with the algal bloom will not be achieved and could additionally lead to high mortality resulting from competition with the holoplankton (Findlay et al., 2009b).

By investigating the effects of pH at a physiological level we gain a greater understanding of how invertebrate internal systems function to cope with the changing environment. Echinoderms are the only invertebrates whose calcified support is endoskeletal, (like chordates, the skeleton is enclosed within at least one epithelial integument (Hofmann et al., 2008)). Other marine invertebrate calcifiers (molluscs, crustaceans, and worms) produce their skeletons as epidermal secretions (exoskeletons) which are in direct contact with their environment (Dupont and Thorndyke, 2009). Calcification involves processes that depend on acid-base status and the maintenance of a suitable microenvironment local to the site of skeletogenic calcification (Dupont and Thorndyke, 2009). Larval survival is crucial to determine recruitment success (Brennand et al., 2010). As the ocean becomes increasingly hypercapnic and seawater pH decreases, the ability of organisms to maintain extracellular pH homeostasis may be compromised, therefore the ability to sufficiently calcify is also compromised. Decreases in larval growth have been linked to a variety of environmental stressors including acidification (Clark et al., 2009), temperature (Byrne et al., 2010b), salinity (Zimmerman and Pechenik, 1991), and pollution (Baussant et al., 2011) in a range of marine invertebrates. Delayed growth is due to decreased energy input (e.g. food

112 intake), by increased energy loss (e.g. respiratory or excretory energy loss) or due to a combination of both (Stumpp et al., 2011b).

During sea urchin larval development, 61% of energy input can be allocated to growth and development after larvae start to feed (Stumpp et al., 2011a). Under elevated ρCO2, if growth is decreased by up to 50%, maintenance becomes increasingly energy consuming under acidified conditions (Stumpp et al., 2011a). In echinoderms food quantity greatly influences survival and growth; feeding is one of the key physiological processes influenced by ocean acidification, with studies showing larval feeding efficiency at low pH is significantly lower than in the control (Dupont and Thorndyke, 2008). This lack of feeding efficiency may compromise calcification and reproduction as a result of the lack of maintenance from internal regulatory systems (Figure 4.1).

Ocean Acidification

OUTPUT (Growth, calcification, INPUT MAINTENANCE (Feeding) (Metabolism, reproduction) respiration)

ECOLOICAL EFFECTS (Predation, competition)

Figure 4.1 Ocean acidification can directly (delays in growth/calcification) or indirectly (longer in the plankton increases predation) affect many physiological processes with consequences on fitness and survival. Modified from Dupont and Thorndyke , 2008 p3131.

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4.3 Climate change and multiple stressors

In addition to ocean acidification, future climate change predictions indicate that marine life forms are threatened by increasing ocean temperature (Harley et al., 2006), sea level rise (Bernstein et al., 2008), and increasing in UV radiation (Harley et al., 2006). Marine invertebrate larvae are small and extremely vulnerable, and in addition, unlike the adult form that are at least partially mobile, the early larval stages are pelagic and have little control over their ambient environment, living in surface waters where stressors such as UV and temperature will have the greatest effect (Leong and Manahan, 1997).

For many organisms, subtle changes in single environmental stressors will not be enough to cause detrimental damage on its own. Multi-factorial studies, such as those by Byrne et al. (2009, 2010 and 2011) indicate that the effects of ocean warming and acidification can act synergistically. In the sea urchin Heliocidaris erythrogramma, development was not affected by pH until a lowering of 0.6 units, yet ocean warming had major negative effects on development. This species may not be able to form a skeleton in a warm ocean, regardless of pH (Byrne et al., 2009). Another study on the adult Antarctic brittle star Ophionotus victoriae indicated that this species was extremely sensitive to small long-term seawater temperature increases, and acclimation to a 2°C temperature increase was not possible (Peck et al., 2009). In the scleractinian coral, Stylophora pistillata, calcification decreased by 50% when temperature and ρCO2 were both elevated, whereas calcification under normal temperature did not change in response to an increase in ρCO2 (Reynaud et al., 2003). In contrast, some studies have shown that warming ameliorates the negative effects of ocean acidification (Brennand et al., 2010, Byrne et al., 2011).

Regulated set points of acid regulation are dynamic depending on the physiological condition of the organism and are influenced by ambient parameters such as temperature and CO2 (Riebesell et al., 2010). Thus including temperature as an additional stressor is more realistic, as global temperatures are predicated to rise 2°C by the end of the century (Bernstein et al., 2008). Increased temperatures are also known to affect physiological processes such as protein synthesis and acid-base balance (Somero, 1995, Whiteley et al., 2001). Tropical species are more heat-tolerant than

114 temperate species, yet are the more threatened by future climate change when compared with species from mid-latitudes because tropical species live closer to their upper thermal tolerance limits and, in some cases, live at temperatures above those at which physiological processes exhibit their thermal optima (Somero, 2010). Additionally, the range of temperatures experienced by an ectotherm may strongly determine its capacity for coping with rising temperatures. Thus, polar ectotherms, especially marine species living at constant near-freezing temperatures, may share with warm-adapted tropical species a high vulnerability to climate change (Somero, 2010).

The responses of our five study species to reduced pH levels during early developmental stages may be exacerbated or perhaps ameliorated by rises in sea temperature. It is important to take into account such realistic scenarios, noting that ocean acidification will not occur on its own. An integrative approach of multiple environmental stressors will provide more reliable results about how species may react to climate change. Responses will be species-specific, with some reacting to pH (Clark et al., 2009, Ericson et al., 2010), some to temperature (Byrne et al., 2009), and some affected only by the coupling of the two (Brennand et al., 2010, Byrne et al., 2011).

4.4 Future research

Laboratory based experiments, such as this study, with controlled parameters (pH, temperature etc.) can provide us with insight into the physiological responses to reduced seawater pH. Any results do, however, need to be interpreted carefully as confounding variables experienced in the natural world are not taken into consideration. These include synergistic environmental factors such as future temperature rises predicted to occur alongside acidification.

In this study I speculate that pHi is being compensated through changes in the pHe. It would have been interesting to use an additional method to measure pHe to validate our HPTS readings, such as a pH electrode, although this technique to measure pHe involves disturbance and stress responses (including shifts in the acid-base status, due to changes in metabolic rate, muscular contraction, release of stress hormones, which could give unrealistic readings of pHe (Pörtner et al., 2010). More ideally, we would use a non-invasive technique such as another pH sensitive dye, BCECF or SNARF,

115 both of which have been used to monitor intracellular pH also (Han and Burgess, 2009). This would then give a better idea of the compensation theory. It would also be valuable to identify limits of compensation inter-specifically and at different life stages

This study did not examine the gastrula or blastula stages of temperate or tropical species, for future studies it would be beneficial to incorporate these stages to get a better idea of how changes in pHe develop. To date, previous studies on ocean acidification responses in echinoderms and development have typically focused on the early period of the larval stage (Clark et al. 2009; Kurihara & Shirayama 2004; Ericson et al. 2010). We know little, however, about the long term effects of pH during later juvenile development, when critical processes such as the development of the rudiment and metamorphosis are occurring. Additionally, acclimation or adaptation may shift the mechanisms and set points of acid–base regulation and may thereby compensate for the

CO2-induced acid–base disturbance and its effect on physiological processes, including calcification (Portner, 2008). Development of a quantitative acid-base stases should include parallel analysis of acid-base parameters in both intra- and extra cellular fluids as well as ambient water (Riebesell et al., 2010).

Future studies also need to address the effects of acid-base variables on metabolic processes and species performances (e.g. growth, shell structure, calcification, reproduction) and fitness (Riebesell et al., 2010). Measurements of metabolic rates should ideally be included as multi-stressor, previous studies have found increased metabolism from increased temperature has reduced the effects of ocean acidification (Brennand et al., 2010, Byrne et al., 2011). This inclusion of metabolism and respiration rates in this study could have provided more details on how metabolism rate might influence pHe.

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4.6 Conclusions

• Morphological development was either delayed or abnormal in all species as seawater pH decreased, with the greatest morphological differences in the lowest treatment (pH 7.4 or pH 7.2) for all species.

• This study, for the first time, shows how pHe can be measured in living echinoderms using the fluorescent probe HPTS.

• The results of this study indicate that extracellular pH (pHe) is affected by an experimental lowering of seawater pH during echinoderm development; - pHe differed among body compartments with the larval gut consistently showing higher pHe than the oesophagus or arms region. - Overall, pHe decreased in each body compartment with decreasing seawater pH.

• The results were species specific and there was latitudinal differences; - Polar species had a naturally lower pHe under ambient conditions, therefore, as seawater pH dropped the pHe dropped and consequently remained below pHe of the temperate and tropical species in most treatments. - S. neumayeri showed a tipping point at treatment pH 7.2 as pHe significantly dropped in all larval regions. - pHe differences among species suggest that physiological capacities to regulate pHe may vary across latitudes, possible due to difference in metabolic rates in polar, temperate and tropical species.

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Appendices .

Appendix 1

The effects lowered seawater has on early cell development. pH 8: predominantly cells are spherical shape and blastomeres are even in size. pH 7.8/7.7: most species cells are still spherical in shape, with regards to slightly abnormal cells in P. huttoni and a slightly unsmooth blastoderm in A. placenta blastula. pH 7.6/7.5: delayed cell development in E. chloroticus, irregular embryos for P. huttoni and S. neumayeri. pH 7.4/7.2: Delayed cell development for E. chloroticus, delayed and abnormal cell development for P. huttoni, abnormal blastoderm development of A. placenta, irregular shaped cells of S. neumayeri, O. validus has irregular blastula development with disrupted cell organisation around the blastoderm.

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a. 9 E vechinus chloroticus y = 10.924x3 - 16.786x2 + 8.7097x + 5.1764 R² = 0.9589 8.5

7.5

6.5

6 Seawater pH Seawater 5.5

5 4.5 4 0 0.1 0.2 0.3 0.4 0.5 0.6 0.7 0.8 0.9 1 exc exc Ratio λ 2/λ 1

b. 9 Psudechinus. huttoni y = 12.52x3 - 18.544x2 + 9.1582x + 5.1608 8.5 R² = 0.9776 8 7.5 7 6.5 6 Seawater pH Seawater 5.5 5 4.5 4 0 0.2 0.4 0.6 0.8 1 exc exc Ratio λ 2/λ 1

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10 c. Arachnoidies. placenta y = 3.1447x3 - 10.119x2 + 11.449x + 2.2453 9 R² = 0.978

Seawater pH Seawater 5

2 0 0.5 1 1.5 2 exc exc Ratio λ 2/λ 1

9 d. Odontaster. validus y = 24.711x3 - 39.175x2 + 20.113x + 2.9342 R² = 0.9813 8

5 Seawater pH Seawater 4

2 0 0.2 0.4 0.6 0.8 1 1.2

exc exc Ratio λ 2/λ 1

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8 e. Sterechinus neumayeri y = 12.046x3 - 19.349x2 + 11.159x + 4.2428 7.5 R² = 0.9625

6.5

5.5

5 Seawater pH Seawater

4.5

3.5

3 0 0.2 0.4 0.6 0.8 1 exc exc Ratio λ 2/λ 1

Appendix 3 One-way ANOVA on the abnormality of early cell development for a) E. chloroticus early b) A. placenta c) P. huttoni , d) S. neumayeri and d) O. to identify significant differences among pH treatments. Embryos were grown in ambient and three lowered seawater pHs.

a) Evechinus chloroticus

Fertilised egg AOV Source DF SS MS F-ratio p pH 3 400 133.33 2.66 0.12 Error 36 400 50 C. Total 39 800 2-cell AOV Source DF SS MS F-ratio p pH 3 12691.66 4230.55 46.15 <0.001 Error 36 733.33 91.66 C. Total 39 13425 4-cell AOV Source DF SS MS F-ratio p pH 3 16400 5466.66 93.71 <0.001 Error 36 466.66 58.33 C. Total 39 16866.67

135 b) Arachnodies placenta blastula AOV Source DF SS MS F-ratio p pH 3 4425 1475 25.29 <0.001 Error 20 466.67 58.33 C. Total 23 4891.67

c) Pseudechinus huttoni

Fertilised egg AOV Source DF SS MS F-ratio p pH 3 8966.67 2988.89 59.78 <0.001 Error 28 400 50 C. Total 31 9366.67 2-cell AOV Source DF SS MS F-ratio p pH 3 4326.91667 1442.30556 86.1077944 <0.001 Error 28 134 16.75 C. Total 31 4460.91667 4-cell AOV Source DF SS MS F-ratio p pH 3 4066.67 1355.56 54.22 <0.001 Error 28 200 25 C. Total 31 4266.67

d) Sterechinus neumayeri

Fertilised egg AOV Source DF SS MS F-ratio p pH 3 2400 800 5.33 0.02 Error 24 1200 150 C. Total 26 3600

e) Odontaster validus

Fertilised egg AOV Source DF SS MS F-ratio p pH 3 300 100 2 0.19265743 Error 36 400 50 C. Total 39 700

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Appendix 4 One-way ANOVA on the morphometric components for a) E. chloroticus early b) A. placenta c) P. huttoni , d) S. neumayeri and d) O. validus to identify significant differences among pH treatments. While similar in developmental stage (4-armed pluteus or gastrula), larvae varied in age.

a) Evechinus chloroticus

Length AOV Source DF SS MS F-ratio p pH 3 8771.14 2923.71 3.13 0.04 Error 36 33631.3 934.20 C. Total 39 42402.44 Width AOV Source DF SS MS F-ratio p pH 3 37743.74 12581.24 3.97 0.02 Error 36 113839.3 3162.20 C. Total 39 151583.042 Rod 1 AOV Source DF SS MS F-ratio p pH 3 16872.83 5624.27 3.83 0.02 Error 36 52840.44 1467.79 C. Total 39 69713.28 Rod 2 AOV Source DF SS MS F-ratio p pH 3 19134.41 6378.13 5.30 <0.01 Error 36 43283.33 1202.31 C. Total 39 62417.75

b) Arachnoides placenta Length AOV Source DF SS MS F-ratio p pH 3 31041.44 10347.14 17.64 <0.001 Error 36 21109.27 586.36 C. Total 39 52150.71 Width AOV Source DF SS MS F-ratio p pH 3 20053.23 6684.41 7.42 <0.001 Error 36 32409.43 900.26 C. Total 39 52462.68 Rod 1 AOV Source DF SS MS F-ratio p pH 3 28997.93 9665.97 6.75 0.00098849 Error 36 51504.44 1430.67 C. Total 39 80502.38 Rod 2 AOV Source DF SS MS F-ratio p

137 pH 3 24849.87 8283.29 6.72 <0.01 Error 36 44325.57 1231.26 C. Total 39 69175.45 c) Pseudechinus huttoni Length AOV Source DF SS MS F-ratio p pH 2 25149.95 12574.97 16.47 <0.001 Error 27 20604.66 763.13 C. Total 29 45754.62 Width AOV Source DF SS MS F-ratio p pH 2 25696.13 12848.06 9.41 <0.01 Error 27 36859.77 1365.17 C. Total 29 62555.90 Rod 1 AOV Source DF SS MS F-ratio p pH 2 13244.77 6622.38 15.37 <0.01 Error 27 11629.55 430.72 C. Total 29 24874.32 Rod 2 AOV Source DF SS MS F-ratio p pH 2 14280.74 7140.37 10.57 <0.01 Error 27 18224 674.96 C. Total 29 32504.74 d) Sterechinus neumayeri Length AOV Source DF SS MS F-ratio p pH 3 1273.4 424.46 2.64 0.06 Error 36 5778.2 160.50 C. Total 39 7051.6 Width AOV Source DF SS MS F-ratio p pH 3 2308.96 769.65 6.65 <0.01 Error 36 4160.4 115.56 C. Total 39 6469.36 Rod 1 AOV Source DF SS MS F-ratio p pH 3 547.82 182.60 4.28 0.01 Error 36 1534.27 42.61 C. Total 39 2082.09 Rod 2 AOV Source DF SS MS F-ratio p pH 3 441.21 147.07 2.46 0.08 Error 36 2150.25 59.72 C. Total 39 2591.46 e) Odontaster validus 15 days Length AOV Source DF SS MS F-ratio p pH 3 4288.27 1429.42 1.83 0.16

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Error 36 28114.98 780.97 C. Total 39 32403.26 Width AOV Source DF SS MS F-ratio p pH 3 16141.53 5380.51 27.16 <0.001 Error 36 7129.75 198.04 C. Total 39 23271.29

Odontaster validus bipinnaria Length AOV Source DF SS MS F-ratio p pH 3 76240.74 25413.58 20.51 <0.001 Error 36 44608.88 1239.13 C. Total 39 120849.63 Width AOV Source DF SS MS F-ratio p pH 3 83168.24 27722.74 21.75 <0.001 Error 36 45884.22 1274.56 C. Total 39 129052.45

Sterechinus neumayeri gastrula Length AOV Source DF SS MS F-ratio p pH 3 1131.48 377.16 7.31 <0.001 Error 36 1856.88 51.58 C. Total 39 2988.37 Width AOV Source DF SS MS F-ratio p pH 3 195.94 65.31 2.30 0.09 Error 36 1019.83 28.32 C. Total 39 1215.78

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