1 Viability of Symbiodinium Dispersed by the Stoplight Parrotfish

Home , Orbicella annularis, Porites porites, Stoplight parrotfish

Viability of Symbiodinium dispersed by the stoplight parrotfish Sparisoma viride

Trigal Magala Velásquez-Rodríguez1, John Mario González2, Juan Armando Sánchez11

Abstract: Herbivorous fishes produce profound effects on the coral reef dynamics and represent a major role as resilience drivers. The finding of viable Symbiodinium cells in the stoplight parrotfish Sparisoma viride faeces supports the notion of these fishes as vectors of a free-living Symbiodinium (FLS) reservoir. However, the magnitude and potential effect of the zooxanthellae dispersed by S. viride was until now unknown. Thus, the purpose of this study was to estimate the FLS viable cells dispersal capacity by S. viride faeces through reefs. We aimed to quantify the number of viable vs. dead cells according to the size faeces pellets. Using a standardized protocol of flow cytometry, we found that to 23,12%±1,50 of the total zooxanthellae are viable in S. viride faeces. Moreover, there are resilience consequences related to S. viride FLS endozoochory evident with the constant input of viable symbiotic zooxanthellae in reefs. All fecal depositions analyzed contained a significant viable cells presence, suggesting a daily functional role that operates in every patch reef composed by S. viride biomass. This outstanding result, confers S. viride similar ecological attributes with herbivorous seeds dispersers in terrestrial ecosystems based on an analog endozoochorous strategy. We propose Symbiodinium icthyochorous dispersal as a new coral reef resilience driver. S. viride conservation efforts focus on overfishing regulation are a priority for the maintenance of its ecological functions and reefs` resilience feedbacks.

Keywords: free-living Symbiodinium, ichthyochory, viability, parrotfishes, Sparisoma viride, reef resilience.

Introduction: Coral reefs are among the most productive and diverse ecosystems in the world (Bonaldo, Hoey, & Bellwood, 2014). The contribution regarding economic, cultural, aesthetic and ecosystem services such as coastal protection, tourism and secure food sources for millions of people, confer coral reefs a unique value as research and conservation targets (Adam, Burkepile, Ruttenberg, & Paddack, 2015; Aswani et al., 2015; FAO, 2014). As a result of synergic natural and anthropogenic threats operating through hundreds of years, coral reefs face the Anthropocene era with high rates of degradation and increasing stress scenarios (Aronson & Precht, 2006; Mora, 2008; Pandolfi, 2015). Therefore, the current and future global challenge is to identify, understand and design strategies for the maintenance of coral reefs ecological functions that confers them resilience and sustainability (Bellwood, Hughes, Folke, & Nystrom, 2004).

A major source of resilience capacity loss is the partial or total decrease of consumer functional groups (Bonaldo, Hoey, & Bellwood, 2014; Plass-Johnson, Ferse, Jompa, Wild, & Teichberg, 2015; Poore et al., 2012). Thus, fishes are considered the main consumers in coral reefs due to their roles in the transfer of energy between trophic groups, benthic algae communities composition, biomass and productivity shape

1 Laboratorio de Biología Molecular Marina, Departamento de Ciencias Biológicas, Universidad de los Andes. 2 Laboratorio de Ciencias Básicas Médicas, Departamento de Medicina, Universidad de los Andes.

1 effect, substrate availability for coral settlement, bioerosion, and sediment transport (Bellwood, Hughes, Folke, & Nystrom, 2004; Bonaldo et al., 2014; Mumby et al., 2006). In a global scale, overfishing constitutes the primary source of ecological functions decrease in reefs (MacNeil et al., 2015). Consequently, different size individuals and species are caught faster than others, generating a selection pressure which alters the base of trophic chains and biogeochemical processes carried by primary producers (Bellwood, Hoey, & Hughes, 2012; Poore et al., 2012).

Massive herbivorous fishes extraction produce profound alterations in the reef dynamic and represent broad concern due to their role as resilience driver (Adam, Burkepile, Ruttenberg, & Paddack, 2015; Mora, 2015). In this sense, herbivorous large-bodied functional groups biomass contribute to control the benthic primary producers, promoting corals settlement, growth, and survivorship in the competitive relationship with algae (Bennett, Wernberg, Harvey, Santana-Garcon, & Saunders, 2015; Edwards et al., 2014).

Parrotfishes (family Labridae, tribe Scarini) are considered the dominant herbivores in the Caribbean (Burkepile et al., 2013; Mumby et al., 2006). This monophyletic group of 10 genera and approximately 100 species with circumtropical distribution is the main contributor to consumer ecological functions in reefs (Bonaldo et al., 2014; Comeros-Raynal et al., 2012). Their beak-like oral jaws (oral teeth fused) and pharyngeal jaw apparatus, constitute functional innovations that conferred specialized feeding morphology making parrotfishes unique within reef fishes. As a result, they effectively grind calcareous material and feed on multiple substrata (algal turfs on dead coral, macroalgae, seagrass and live corals) depending on their three functional groups (Bonaldo et al., 2014; Price et al., 2010). In this sense, browsers consume macroalgae without leaving scars on the substratum and contribute to prevent coral-macroalgae phase shifts. In contrast, scrapers bite slightly into the substrate leaving a scrape signal of the removal material, while excavators make prominent scars as a result of deep bites during feeding (Francini-filho, Moura, Ferreira, & Coni, 2008; Sánchez, Gil, Chasqui, & Alvarado, 2004).

Among parrotfishes ecological functions, corallivory represents a highly specialized one related with excavators feeding mode (Bonaldo et al., 2014). The stoplight parrotfish Sparisoma viride (up to 510mm fork length terminal phase) is the primary excavator at the Colombian Caribbean (Francini-filho et al., 2008; Reyes-Nivia, Garzón-Ferreira, & Rodríguez-Ramírez, 2004; Sánchez et al., 2004). For S. viride, live coral represents near 1% of the total diet, being its primary feeding substratum the Epilithic Algal Matrix (EAM) on dead coral surfaces (McAfee & Morgan, 1996), which is a complex aggregation of short filamentous algae growing in turf, mixed with microalgae, macroalgae propagules, sediment, detritus and fauna associated (Bonaldo et al., 2014). Regarding coral species preference, S. viride forages mainly on Orbicella annularis, Montastrea cavernosa, O. faveolata, O. franksi, Agaricia agaricites, Porites porites, P. astreoides, Pseudodiploria strigosa and Colpophyllia natans (Bonaldo et al., 2014; Castro-Sanguino, 2010; Reyes-Nivia et al., 2004; Rotjan & Lewis, 2005; Sánchez et al., 2004).

During corallivory, S. viride ingests coral fragments that pass through the digestive tract and are released as fine sediment in subsequent defecation (Perry, Kench, O’Leary, Morgan, & Januchowski-Hartley, 2015). One functional ecology key finding made by Porto et al. (2008) and Castro-sanguino & Sánchez (2012) is the ability of unicellular dinoflagellate endosymbionts cells, referred as zooxanthellae (Symbiodinium), to survive S. viride gut passage and maintain viable in faeces. Symbiodinium studies are

2 numerous due to their role as cnidarian symbionts in the holobiont coevolution processes, specifically in scleractinian corals (Levin, Suggett, Nitschke, van Oppen, & Steinberg, 2017). The mutualistic relationship Symbiodinium-coral is based on the exchange of algal photosynthetic products and animal nitrogen compounds discarded. In detail, Symbiodinium translocates photosynthetically-fixed carbon to host as energy demand supplement promoting its calcification rates; and complementary, host confers to symbionts the inorganic nutrients required for photosynthesis and offers protection from predators (Muller-Parker, D’Elia, & Cook, 2015; Stambler, 2011).

The occurrence of viable Symbiodinium cells in S. viride faeces means this parrotfish acts as a key dispersal agent (endozoochory) and supports the maintenance of zooxanthellae environmental reservoirs in reefs (Castro-sanguino & Sánchez, 2012; Porto et al., 2008). Free-living Symbiodinium (FLS) are ubiquitous in environmental reservoirs and refers to cells living outside a cnidarian host that conserve the capacity of establishing symbioses (Thornhill, Howells, Wham, Steury, & Santos, 2017). Particularly, FLS possess a motile stage with two flagella (dinomastigote). This swimming stage often alternates with the zooxanthellae coccoid one according to a diurnal pattern registered in cultures (Muller-Parker et al., 2015). The links between FLS and zooxanthellae in symbiosis are: firstly, FLS were symbiotic zooxanthellae before a host expelled it; secondly, FLS are dispersed by host larvae or corallivorous fishes; and thirdly, FLS are available for juveniles or adult hosts uptake (Cunning, Yost, Guarinello, Putnam, & Gates, 2015).

Understanding FLS physiology, reproduction, distribution, dispersion and population genetics is currently limited and a priority as part of Symbiodinium life cycle comprehension (M. R. Nitschke et al., 2016; Porto et al., 2008). Research efforts developed in order to identify environmental reservoirs confirmed Symbiodinium presence in the water column (Granados-Cifuentes, Neigel, Leberg, & Rodriguez-Lanetty, 2015; Littman, van Oppen, & Willis, 2008; Takabayashi, Adams, Pochon, & Gates, 2012); benthic macroalgae beds (Porto et al., 2008; Venera-Ponton, Diaz-Pulido, Rodriguez-Lanetty, & Hoegh-Guldberg, 2010) and sediments (Granados-Cifuentes et al., 2015; Littman et al., 2008; M. R. Nitschke et al., 2016; Takabayashi et al., 2012). Additionally, respect to other corallivorous fishes as FLS dispersers, Muller- Parker (1984) registered Arothron meleagris, Chaetodon auriga and C. unimaculatus releasing fecal pellets composed basically by photosynthetically active zooxanthellae from the predated sea anemone Aiptasia pulchella.

The current compiled evidence about S. viride FLS dispersion includes, first the confirmation by light microscopy and enumeration of Symbiodinium in fresh faeces, reporting densities from 73 to 38.3x103 cells ml-1 (Castro-sanguino & Sánchez, 2012). Second, successful isolation and culture of Symbiodinium- like cells from faeces enrichment in zooxanthellae selective medium (f/2) (Castro-sanguino & Sánchez, 2012; J., 2007; Porto et al., 2008). Third, molecular identification of Symbiodinium clades from fresh faeces and culture cells, reporting clades A, B, and G using the chloroplast ribosomal marker (cp23S- HVR) for Colombian Caribbean localities (Castro-sanguino & Sánchez, 2012). Finally, S. viride dispersion potential estimation of FLS in terms of movement through territory areas; thus, Castro- Sanguino & Sánchez (2012) demonstrated zooxanthellae transport at a local level, reporting 35m as the maximum distance between faeces ejected sites.

3 In the knowledge built up to date, FLS viability dispersed by S. viride faeces has been inferred indirectly from two methods: cell identification of reproductive stages (coccoid, motile and mitotic forms) associated with asexual reproduction viability; and cell capacity to increase density and grow in cultures (Buysschaert, Byloos, Leys, Van Houdt, & Boon, 2016; Castro-sanguino & Sánchez, 2012). Complementary, Delgado (2013) approached to the zooxanthellae dispersed viability estimation using SYSTOX ® Green, finding up to 15.78% viable cells/individual; however, she emphasized a high non- explained variation between samples viability values. In this context, additional efforts to clarify the viability of FLS dispersed by S. viride faeces are a priority. Similar studies have estimated the zooxanthellae viability in Dermasterias imbricate seastar, Clinocottus globiceps fish and Aeolidia papillosa nudibranch faeces, which predate the anemone Anthlopeura elegantissima. As main conclusions, Seavy & Muller-Parker (2002) and Bachman & Muller-Parker (2007) confirmed the release of viable zooxanthellae based on the mitotic index, productivity and photosynthetic pigments indicators, demonstrating similar productivity, lower photosynthesis rates, and no significant division process differences, in contrast to Anthlopeura elegantissima freshly isolated zooxanthellae. Finally, Augustine & Muller-Parker (1998) reported Symbiodinium cells digestion and non-viable cells presence.

Cell viability is a key variable in endozoochory studies, due to its ecological implications as effective dispersal mechanism (Wilkinson, Lovas-Kiss, Callaghan, & Green, 2017). Specifically, a cell viability assay differentiates between viable cells (living and healthy), and non-viable cells (dead or degrading). As a result, a viability value is estimated based on the percentage of live cells respect to the total cells population number (live and dead) (Burtscher, May, Downs, & Bartlett, 2015; Buysschaert et al., 2016). In general, Symbiodinium viable possess coccoid shape with intact cell membrane, spherical nucleus, well- developed elongated chloroplasts and defined pyrenoid bodies. Conversely, non-viable Symbiodinium present unrecognizable organelles (cytoplasmic ruptures), and cell membrane fragmented or intact cell membrane without defined cytoplasmatic material (M. Nitschke, 2015; Strychar, Coates, Sammarco, Piva, & Scott, 2005). Cell viability assays conclude about the capacity of cells populations of sustaining life (Burtscher et al., 2015).

New arguments about FLS viability in S. viride faeces are crucial to understanding in situ potential ecological impacts. Thus, the purpose of this study was to estimate the FLS viable cells dispersal capacity by S. viride faeces through reefs. We aimed to quantify the number of viable vs. dead cells according to the size faeces pellets. Our hypothesis was: there are no significant differences between the viable and non-viable percentage of cells dispersed by S. viride faeces. This work provides evidence about the understanding of a new S. viride functional role that confers reefs resilience. Conservation efforts must be conducted to maintain this S. viride function through time in the current challenge of reefs sustainability.

Methods:

Study Area The fieldwork was carried out in the Archipelago of San Andrés, Old Providence and Santa Catalina (Southwestern Caribbean), which is composed of oceanic islands, submerged banks and atolls located approximately 800 km northwest of Colombia (Puyana, Acosta, Bernal-Sotelo, Velásquez-Rodríguez, & Freddy, 2015). This territory is part of the Seaflower Biosphere Reserve declared by UNESCO in 2005,

4 and considered the largest Marine Protected Area (MPA) in the Caribbean (Castaño-Isaza, Newball, Roach, & Lau, 2015). Seaflower MPA is especially global relevant due to contains over 200.000 ha of productive open-ocean reefs and extended reefs associated with diverse ecosystems (The CaMPAM Mapping Project, 2017), whereby is known as the third largest coral reef barrier worldwide (Baine, Howard, Kerr, Edgar, & Toral, 2007). Specifically, this study was developed in San Andrés Island, characterized by 15 km of long reef barrier presence (The CaMPAM Mapping Project, 2017). The sampling was conducted at Nirvana Diving Site in 12° 30′ 05″N and 81° 43′ 56″ W coordinates.

Figure 1. Map of San Andrés Island, showing the sampling site.

Faeces sampling Observational samplings of focal male adult individuals (terminal phase) of S. viride were conducted using SCUBA diving in a total area of 5.000m2, constituted by different patches reefs (depth 3 to 12m). The standardized time for focal observations was 10 minutes per individual; preferred individuals were those foraging near to the bottom, due to the consistent quality of the faeces released. During this time,

5 one pellet faeces sample per individual was collected immediately after released on substrate using 10 ml syringes. A total of 65 pellet faeces samples were collected. Additionally, a sediment control sample was taken (60 cm horizontally to the pellet faeces fall place). Each syringe was disposed in a Ziploc bag marked with the sample type and safely kept. Generally, pellet faeces were collected on sediment bottom; and occasionally on live and dead coral surfaces. After the immersion, samples were transfer to 15ml falcon tubes and formalin 10% was added to content fixation. This sampling was developed during june/2016. Underwater time corresponded to 24 diving hours.

Faeces filtration In the laboratory, each sample was filtered using a plastic Büchner funnel coupled with a vacuum pump and a previously weight led Whatman filter paper grade 4 (25-30um mesh size). Sample scattered on the filter paper was washed with 0.22 um Filtered Sea Water (FSW) until recovering a final volume of 50 ml. After, the filter was dried at room temperature, and its new weight with the sample sediment retained on it was registered. Zooxanthellae cells size range from 5 to 20 um (Muller-Parker et al., 2015), thus were expected to be in the filtrate recovered. Each 50 ml falcon tube was centrifuged at 3100 rpm during 5 min. Subsequently, the pellet was re-concentrated on 250 ul of FSW (from here considered as the sample). The difference between blank filter paper and dried sediment sample on filter paper was considered as the faeces dry sediment miligrams collected. This process was applied to the control samples too. Filtered samples were protected of light incidence, due to the possible autofluorescence degradation of zooxanthellae.

Microscopy verification and enumeration Microscopic observation to verify Symbiodinium cells presence in each sample was carried out on an Improved Neubauer haemocytometer (Hausser Scientific, USA), under a compound light microscope Olympus CX21, using 40X. Additionally, a subsample (50 ul) of 30 S. viride concentrated samples randomly selected was used for enumeration and cell concentration report. A total of 4 counts were done per sample. Control samples were counted following the same steps. Symbiodinium recognition was based on previous observations of tissue samples and zooxanthellae extruded from Pterogorgia guadalupensis, and from published identification photos on scientific papers. Thus, cells were identified using characters as unicellular circular shape, golden-brownish color, elongated and reticulated chloroplast and large pyrenoid body (Nitschke, 2015).

Viability quantification by flow cytometry FCM Standardization Quantification of viable and dead Symbiodinium cells in S. viride faeces samples was carried out by flow cytometry (FCM). This automated method represents an alternative to traditional microscopy techniques, which are related to time-consuming samples processing, subjectivity in observations and lowly reproducibility count to count (Lee, Sheng, Yeo, & Sin, 2012). FCM identifies cells populations based on: first, particles light scattering principles, and second the excitation and emission of fluorescence by particles fluorochromes; both as the response to a light beam (Krediet et al., 2015).

In detail, a cell solution enters into the sample injection probe, in which there is a stable stream (of generally PBS solution) that obligates each particle to move one by one. During the travel, each particle passes through a flow cell, the site in which is stimulated using laser illumination (488nm). As a result, the

6 particle emits a response expressed in two measures of light scattering that are detected and converted into electrical signals by a light detector. These measures emitted by particles are digitalized as: first, the forward scatter (FSC) or the signal related to the cell size and shape; and second, the side scatter (SSC) or the signal indicative of cell granularity. Based on FSC and FSC parameters are specific to each particle type, FCM identifies cells population composed of the same particles types (McCarthy & Macey, 2007).

Additionally, the FCM use the fluorescence emission by particles as identification criterion. After the laser beam crosses particles, excites their fluorochromes and light is emitted at different wavelengths. These wavelengths are split using optical filters in specific colors and detected by optical detectors known as photomultiplier tubes (PMT). As a result, the PMT generate an electrical pulse converted into readable signals by computer software. Those signals are expressed as cell events of a population in a dot plot (Papandreou, 2013).

For FCM standardization purposes, preliminary assays were developed using two different samples types; first P. guadalupensis extract from live tissue, and second, P. guadalupensis zooxanthellae cultures aliquots. In the beginning, the process consisted in differentiate cellular components from debris particles based on FSC and SSC. Both parameters indicated heterogeneous size of cellular components in samples.

In the next stage, cellular components were distinguished using fluorescence emission. For this, propidium iodide staining was performed to both types of samples as a way of promoting fluorescence resolution in cell viability quantifications; however, there were not observed significative differences in particles fluorescence, in comparison to non-stained samples. Consequently, was concluded that Symbiodinium autofluorescence signal was pertinent as viability quantification criterion. In this way, a red fluorescence threshold set (670nm long-pass), related to Symbiodinium autofluorescence was established after testing the efficiency of three different PMT (520nm long-pass, 590nm long-pass and 670nm long-pass in cells differentiation. Finally, cells populations in faeces and sediment samples were determined using common FCS, SSC, and fluorescence parameters between particles.

Subsequently, sorting was carried out on the individual defined populations. After, each population sorted was observed using an Improved Neubauer haemocytometer and photographic registers were taken. Additionally, fluorescence microscopy observations were done to corroborate light microscopy findings. Based on the sorted population results, the FCM settings were considered standardized for the S. viride faeces viability quantification.

FCM conditions for viability quantification S. viride faeces samples and sediment controls filtered were analyzed using a BD FACSCanto IITM cytometer, coupled with the FACDS Diva 6.1. Software. A sample volume of 200 ul was used for each run, and a total of 1500-2000 cells were registered. FSC and SSC were collected using a 488nm band-pass filter, with blue light of origin. Additionally, Symbiodinium red autofluorescence signals (chlorophyll a) were collected via 670nm band-pass filter. As a conclusion, FSC and SSC, in combination with the Symbiodinium endogenous autofluorescence were used to discriminate zooxanthellae population from macroalgae fragments, non-algal cells and high quantity of debris in S. viride faeces. The same parameters functioned for zooxanthellae viable and non-viable cells differentiation. Normalization of data was made for assuring an accurate comparison of cell size, fluorescence patterns and viability percentages between

7 populations in the total samples. Table 1 contains the technical details used for the FCM analysis.

Table 1. FCM parameters for Symbiodinium viability estimation.

Logaritmic Parameters Voltage criterion

FSC 210 Yes SSC 222 Yes FITC 560 Yes PE 500 Yes PE-Cy7 600 Yes

Statistical Analysis Viability quantification data were analyzed for normality. Multifactorial ANOVA was used for examining significance of Symbiodinium viability vs. non-viability in S. viride faeces, and in contrast to sediment controls. Additionally, the Person Correlation Coefficient was calculated for testing the relationship between viable and non-viable Symbiodinium populations vs. miligrames of faeces dispersed.

Results:

FCM Method During the standardization process, four cell populations were determined (figure 2): P1 (red), P2 (green), P3 (blue) and P4 (black) in a dot plot. According to population distribution, P2 was identified as the one characterized by higher fluorescence signal emission, size and granular cells content. Besides, P3 showed a lower fluorescence signal than P2, and granular cells too; P1 presented the lowest fluorescence signal vs. P2 and P3; and P4 exhibited non-cellular components or debris.

Figure 2. Populations determined during FCM standardization process.

8 Complementary, sorting results conduced to the four populations identities validation (figure 3). Specifically, viable/healthy Symbiodinium cells were found in the P2 population, and degrading/dead Symbiodinium cells were registered in P3. On the other hand, none Symbiodinium cells were identified in P1 and P4, in which macroalgae fragments and unspecific subproducts were observed in accordance with S. viride diet.

A. B. C. D. P2 P1 P4 P3

P3 P1 P2 P4 P4

Figure 3. Sorted population identities validation: A. P2 contains Symbiodinium with viable/healthy appeareance; B. P3 presents Symbiodinium with degraded/dead characteristics; C. P1 as well as D. P4 does not contained Symbiodinium-like cells.

Additionally, fluorescence microscopy observations to sorted populations were coherent with the previous light microscopy findings. In this sense, P2 contained Symbiodinium cells that presented the highest fluorescence emission; P3 presented zooxanthellae cells with less fluorescence emission in contrast to P2; and finally, P1 and P4 components emitted the lesser fluorescence (figure 4). Light and fluorescence observations in sorted populations supported the validation of FCM standardized parameters.

A B

P2 P3

PP2 P3

Figure 4. Light vs. fluorescence microscopy registers to sorted cells populations (40x): A. P2 Symbiodinium cell emitted higher fluorescence than P3. B. P3 Symbiodinium cell with its characteristic fluorescence.

9 Zooxanthellae verification and enumeration S. viride faeces and control sediment samples were observed under light microscopy to register Symbiodinium presence (figure 5). Zooxanthellae cells were confirmed in 100% of the samples and controls collected. In all cases, cells were observed coccoid in shape with 11-13 um of diameter. Dividing cells stages were evidenced in faeces samples. Intracellular composition patterns varied from well- developed and elongated chloroplasts and defined pyrenoid body to few recognizable organelles presence. Additionally, the color pattern observed corresponded to golden brownish cells to translucent appearance.

Figure 5. Symbiodinium cell identified in S. viride faeces (40x).

Symbiodinium cells density was higher in S. viride faeces than in sediment samples corresponding to 33x102cells 370mg-1 and 11x102cells 240mg-1, respectively. On average, faeces samples registered a dry mass of 370 mg±0,03 and sediment controls presented a value of 240 mg±0,03. Density counts were normalized to compare information between samples, using the mass data registered for each case. Thus, zooxanthellae density was expressed as the number of cells present in 1 mg of dry mass sample collected (figure 6).

8 dry mass) dry -1

2 mgZooxanthellae (cell

0 S. viride Faeces Sediment Control

Figure 6. Symbiodinium density in S. viride faces vs. sediment control.

10 Viability quantification by flow cytometry Accordingly to FCM standardized settings, the four populations validated were identified in all S. viride faeces and sediment controls (figure 7). An entire zooxanthellae population per sample was defined as the sum of P2 (viable/healthy zooxanthellae) and P3 (dead/degrading zooxanthellae) events. Thus, 22855 viable events were registered in P2 vs. 7500 viable ones in P3. Regarding faeces or sediment control mg-1, a total of 62 viable events mg-1 corresponding to P2 were detected, and in contrast, 32 viable events mg-1 in P3 were evidenced. This fact means a proportion of 2 viable cells events registered in faeces samples to 1 viable cell event in sediment control. Conversely, P1 and P4 data were not included in viability analyses due to the absence of zooxanthellae in them. In this context, Symbiodinium viability calculation per sample was estimated based on the relation between the specific live cells frequency and the total cells population frequency (live and dead), expressed as a percentage. The same calculation was done for non- viable percentage estimation.

A. B.

Figure 7. FCM dot plots registering the four populations validated during standardization in: A. S. viride faeces, and B. sediment control samples. Viable Symbiodinium cells were observed in P2; and non-viable in P3 in both types of samples.

Symbiodinium viability in S. viride faeces corresponded to 23,12%±1,50, whereas its non-viability to 76,88%±1,50 (figure 8). In another hand, 20,02%±5,36 of Symbiodinium cells found in sediment control samples showed viability, and 79,98%±5,36 exhibited non-viability. Additionally, viable and non-viable percentages registered for S. viride faeces were significantly different (F=481,2, p<0,001); this pattern was confirmed in sediment controls too. In contrast, no significant differences were registered between Symbiodinium viability in S. viride faeces vs. sediment control samples (F=0,1, p>0,7). Finally, non- correlation was observed between the viability percentages obtained to S. viride faeces or control samples vs. the dry mass collected (R-squared=0,023, F=1,067, p>0,307), this means the sample´s mass did not explain the zooxanthellae viability values.

11 79,98 76,88 100

80 20,02

60 23,12

Percentage (%)

0 Viable Non-Viable Viable Non-Viable Cells Cells Cells Cells

S. viride faeces Sediment

Figure 8. Boxplot showing: significant differences between Symbiodinium viability percentages in S. viride samples, and the same tendency in sediment control samples. No significant differences were found in S. viride faeces viability percentages vs. sediment controls.

Discussion: A major resilience role can be attributed to S. viride as a functional ecological feedback of FLS endozoochory on reefs. We confirmed Symbiodinium presence in 100% of S. viride faeces sampled, in which an average density of 33x102cells 370mg-1 dry weight faeces was enumerated. From these cells, the 23,12%±1,50 is able to maintain viability after the digestion process through S. viride gut. Faeces size had no effect on the number of viable FLS quantified. Our results suggest a daily functional role that operates in every reef patch composed by S. viride biomass. This outstanding dispersal capability confers S. viride comparable ecological attributes with herbivorous seed dispersers in terrestrial ecosystems based on the same endozoochorous strategy.

Endozoochory represents the principal seed dispersal strategy in forests (Guy-Haim, Hyams-Kaphzan, Yeruham, Almogi-Labin, & Carlton, 2017). According to endozoochorous ecological principles, the dispersal success depends strictly on the effective release of viable seeds through the landscape (Blanco et al., 2016). This fact is linked to the abundance and distribution of faeces containing viable seeds (Suárez- Esteban, Delibes, & Fedriani, 2013). In the S. viride FLS endozoochory, viable Symbiodinium cells are released through reefs at defecation rates of approximately one pellet 3 min-1 per individual during its higher faeces release hours (10:00-15:00) (Vermeij et al., 2013). Regarding spatial distribution, S. viride cells input occurs in territories of approximately 100 to 300 m2 (medium sized), and in a daily feeding range of 50 to 800 m2 (Bruggemann, Van Kessel, Van Rooij, & Breeman, 1996; Tolimieri, 1998). In this

12 area, S. viride releases faeces into new sites constantly on the move (Sazima, Bonaldo, Krajewski, & Sazima, 2005), and a maximum distance of 35 m exists between them (Castro-sanguino & Sánchez, 2012). Another essential aspect that promotes faeces distribution is their dissolution pattern in the water column. Thus, S. viride faeces usually remain as a solid pellet for a short time after defecation (Goatley & Bellwood, 2010), due to disaggregation by water waves and currents (Hopkinson, 1985). Additionally, this species tend to produce clouds of faeces in response to behavioral patterns, which disperse quickly in the water column (Sazima et al., 2005). These distribution patterns contrast to those of holothurian species, in which faeces disintegrate completely in a time lapse of 8 hours (Isostichopus badionotus), and even remain on the sea bottom for up to 25 hours after deposition (Holothuria mexicana). Pellet dissaggregation is a crucial aspect for particles and nutrients recycling (Conde, Diaz, & Sambrano, 1991), and indeed for FLS distribution on reefs.

Dispersal capability in endozoochorous studies is estimated using data as the density of cells dispersed, defecation rates and viability after release (Vermeij et al., 2013). Based on our density and viability results it is possible to estimate the S. viride FLS dispersion potential. In a conservative perspective, if an individual defecates a 130 mg dry weight faeces pellet (minimal value we registered) each 3 min (rate during higher faeces release hours), then it can disperse >1170 FLS viable cells per day through reefs. Moreover, our results showed that on average the faeces pellets dry weight is 370 mg. In accordance to this, S. viride viable FLS daily dispersal capacity could be as many as 76.000 cells individual-1. Our FLS dispersal estimation is based on Vermeij et al. (2013) defecation rates and could change according to other S. viride populations with local foraging ecological patterns (Castro-Sanguino, 2010). However, our values reported are conservative as we consider the hourly interval with higher defecation frequency exclusively. A higher viable FLS contribution could be occurring in reefs based on a complete daily defecation rates data analysis.

Animal faeces as live zooxanthellae reservoirs have been studied for more than 25 years. However, FLS presence in S. viride faeces study is special in contrast to others cases. First, the majority of literature has confirmed zooxanthellae in animal faeces that establish symbiosis. In this case, the release in faeces constitutes a zooxanthellae density regulation mechanism by the host. Examples are nudibranchs species Dotodoerga paulinae, D. rosea, Catriona maua, Cuthona caerulea, C. granosa, Aelidiella aldieri, Spurilla neapolitana, Melibe engeli and Berghia verrucicornis, in which brownish faeces containing large numbers of undigested Symbiodinium cells were evidenced (Burghardt & Wägele, 2014; Kempf, 1991; Marin & Ros, 1991). Although these zooxanthellae release is not endozoochory (a vector is absent), it is relevant to analyze the cells input on reefs by the mentioned species' faeces in contrast to our findings.

In this sense, giant clams species Tridacna derasa and Tridacna gigas release 4.9x105 cells day-1 and 4.7x105 cells day-1 of intact zooxanthellae in faeces, respectively (Maruyama & Heslinga, 1991; Neo, Eckman, Vicentuan, Teo, & Todd, 2015). These numbers correspond to 11% of the remaining viable cells after digestion due to clams digest approximately 64 to 89% of total zooxanthellae. Additionally, swimming zooxanthellae have been mentioned to occur in their faeces, but in no-significant numbers (Maruyama & Heslinga, 1991). Those motile cells have been reported mainly at the end of the light period during the light and dark cycle (Fitt & Trench, 1983), which represents a limited time in Symbiodinium life cycle. In our results, motile cells were not observed in S. viride faeces due to samples fixation, but this would be interesting to observe; if S. viride could disperse motile zooxanthellae in reefs, a plus in its

13 endozoochory role could be occurring. According to Fitt & Trench (1983), the average swimming speed of motile zooxanthellae is near to 365 µm s-1; this means a swimming pattern of 4 m in 3 h that could be up to 10 m in the daily motility period (8 h). In this context, although these species release a huge quantity of cells vs. S. viride, their resilience effect could be limited by a restricted distribution according to life cycle characteristics (low dispersal capacity and sessile stage in giant clams adults), and a low diversity of Symbiodinium clades in faeces related to species-specific symbiosis.

The second set of evidence about animal faeces containing live zooxanthellae is related to cnidarian predators. Precisely, the gastropod species Coralliophilla meyendorfii, the seastar Dermasterias imbricata and corallivorous fishes as Clinocottus globiceps, Arothron meleagris, Chaetodon auriga and C. unimaculatus constitute endozoochory examples in coral reefs (Augustine & Muller-Parker, 1998; Muller- Parker, 1984; Poulicek, Roberty, Plaza, & Ladriere, 2010). A common characteristic identified in the mentioned studies is the ingestion of cnidarians as the main component of the daily diet during experiments. For example, Poulicek et al. (2010) fed in vitro to Coralliophilla meyendorfii with Anemonia viridis exclusively; and after one week tested the zooxanthellae presence and viability in faeces. Additionally, Muller-Parker (1984) fed collected individuals of Clinocottus globiceps, Arothron meleagris, Chaetodon auriga and C. unimaculatus exclusively with Aiptasia pulchella during 24 hours and after confirmed zooxanthellae in faeces. In this context, our results are the first in testing Symbiodinium presence and viability in faeces obtained directly in the field, as a consequence of natural feeding ecological conditions in a marine vertebrate disperser. Our methodological approach in faeces sampling corresponds to the same used in the majority of terrestrial and freshwater endozoochory studies. Thus, S. viride´s role is comparable to those developed by dispersal vectors in these ecosystems.

Based on the previous arguments, the elucidated capability of S. viride in viable FLS dispersal can be considered a significant functional ecological role in reefs. Consequently, positive ecosystemic feedbacks emerge as benefits linked with the efficient occurrence of endozoochory. Examples of these positive impacts are the dispersal of population with restricted distribution, genetic connectivity maintenance, genotypic diversity increase, and resilience contribution regarding regeneration and restoration after disturbances (Castro-sanguino & Sánchez, 2012; Pollux, 2011; Shikang, Fuqin, & Yuehua, 2015; Tol et al., 2017). In the S. viride role of repopulating FLS environmental reservoirs with viable cells, one outstanding feedback would be the ability of dispersed zooxanthellae to be uptaked by aposymbiotic models (Castro-sanguino & Sánchez, 2012). As Muller-Parker (1984) tested with the FLS released by cnidarian predators and Nitschscke et al. (2016) with FLS on sediments, the zooxanthellae uptake suggests resilience potential in reefs during post-bleaching and other disease recovering processes.

Endozoochory is more than an ecological issue. In this sense, a relevant shaping co-evolution process between disperses and cells dispersed have been occurring at least during 80 Myr. Thus, interactions have been reported between 331 vertebrate species: birds (232), mammals (90), fishes (5), amphibians (1) and reptiles (3), and more than 700 plant species (Bello et al., 2017; De Vega, Arista, Ortiz, Herrera, & Talavera, 2011; Pakeman, Digneffe, & Small, 2002). As a consequence, rapid traits evolution in dispersed species (i.e., seed size change), and even population genetic disruption have been reported (Bello et al., 2017; Pollux, 2011). Based on this terrestrial ecology framework, the FLS dispersal role by S. viride must be understood as a potential driver of Symbiodinium composition structure in reefs, that permeates FLS biology, genetics, and evolution (Pollux, 2011).

According to this ecological and evolutive perspective, it is relevant to analyze the interaction between viable Symbiodinium cells and S. viride. Thus, some pertinent questions to explain our viability results are: What characteristics of S. viride are related to the survivorship of Symbiodinium cells? And what traits of Symbiodinium cells determine their ability to survive and remain viable after the gut passage? With respect to the first question, Augustine & Muller-Parker (1998) reported non-viable zooxanthellae cells in Clinocottus globiceps faeces, and explained this result as the gastric acid action in the fish stomach, which conduce to zooxanthellae cells structural components lysis. In contrast, S. viride possesses a particular digestive tract morphology, which supports our viability results. In detail, its digestive tract is constituted by oral fused teeth, pharyngeal jaws and a long intestine, but stomach and pyloric caeca are absent (Papoutsoglou & Lyndon, 2006). This fact means acidity activity is not present as a mechanism of digesting food. This corporal plan corresponds to that characteristic in herbivorous fishes, who are specialized in grinding mechanisms for breaking ingested material (Lobel, 1981). As a consequence, there are no low pH conditions in the S. viride digestive tract that lyse Symbiodinium cell walls. Of course, grinding through two pairs of jaws represents a high mechanical action due to calcium carbonate trituration, whereby the 76,88%±1,50 of cells released in faeces were found to be non-viable.

On the other hand, our results reported FLS presence in 100% of faeces samples. This means Symbiodinium possesses traits that promote their ubiquity in S. viride faeces, and resistance to the pass through the pharyngeal jaws and long gut. In terrestrial ecology, ubiquitous dispersed seeds share common characteristics as size and shape. Specifically, more abundant recovered seeds in faeces are small and round, traits that correspond to Symbiodinium morphology. Based on Staniforth & Caves (1977) and Thompson et al. (1993), a small spherical morphology skips easily to grinding and gut passage, in contrast to large and elongated cells. As a result these autors predicted a greater potential in dispersal distance. In this sense, spherical morphology of Symbiodinium cells could explain their viability percentage in faeces.

The second trait of Symbiodinium that confers them survivorship during S. viride endozoochory is its cell structure and composition. The characteristics in hardness or toughness in a seed are considered as the most relevant in terms of interspecific viability and survival differences between seeds (Pollux, 2011). As a dinoflagellate, Symbiodinium presents a cell cover called amphiesma, which is composed of a complex arrangement of five membranous layers with internal material (Pozdnyakov & Skarlato, 2012). Also, the cell has a multilobed chloroplast that as a particular trait possesses an envelope made by three different layers (Blank, 1987). Based on this, we attribute the preserved fluorescence signal emitted by zooxanthellae during our FCM application to the chloroplast complex structure, and in general to its external cell protection. Thus, Symbiodinium cell configuration protects it from fatal damage against the high mechanical action of pharyngeal jaws after S. viride ingestion.

Moreover, contrasting our viability estimations to the post-dispersal analysis of released seeds suggests high variability in multiple interactions corresponding to particular life history and traits of species (seed released and disperser). These interactions could explain patterns as our result of a non-faeces-size effect on the density of viable FLS dispersed. Specifically in viability, for example, Pakeman et al. (1999) reported a 27% of viable dispersed seeds in grazeland systems foraged by rabbits, considering it as a significant effect on restoration. In contrast, Blanco et al. (2016) found viabilities from 0% up to 92,7%

15 depending on the plants species dispersed by parrots faeces; and other studies informed intermediate viability values. The important aspect is that in all the cases viability is related to resilience in ecosystems.

Furthermore, control sediment samples showed a similar viability percentages tendency, in comparison to faeces samples. However, a clear difference in zooxanthellae density between sample type was obtained, demonstrating a higher input of FLS through S. viride faeces. As Thornhill et al. (2017) mentioned FLS in reefs are common outside hosts, higher in cell density than those in the water column, and primarily demersal. The potential of those zooxanthellae regarding reservoirs of viable cells has been well documented. Studies have grown cells from sediments and conducted experiments of infection with aposymbiotic larvae and coral recruits (Granados-Cifuentes, Neigel, Leberg, & Rodriguez-Lanetty, 2015; Littman, van Oppen, & Willis, 2008; M. R. Nitschke, Davy, & Ward, 2016; Takabayashi, Adams, Pochon, & Gates, 2012). Consequently, cnidarian models tested have acquired multiple Symbiodinium clades after the exposure to sediments, and its infection capability has been demonstrated. In this way, sediments represents a site in which converge short life FLS from multiple sources, whose function has been proposed as to infect new hosts (Thornhill, Howells, Wham, Steury, & Santos, 2017). Finally, the similar pattern of S. viride and sediment controls viability percentages evidenced could be understood through the link between parrotfish and coral sediment production. According to Perry et al. 2015 (2015), excavators parrotfishes were the major coral sediment producers in their study area, thus FLS in faeces could be “fertilizing” sediments daily with their viable cells.

Our results provide new insights about S. viride as primary components of reefs. Moreover, the known functional roles in herbivory, sediment production, and bioerosion; the role of FLS endozoochory strengthens S. viride relevance in ecosystem dynamic as resilience driver. The challenge now is to understand how to maintain this essential functional role in the current context of reefs at the Caribbean. As Bonaldo et al. (2014) mentioned, parrotfish have been crucial components of the ecosystem dynamics in the past, and most probably will be key in the future shaping of coral reefs.

Conclusions: A major resilience role is attributed to S. viride as an endozoochorous vector of FLS viable cells on reefs. This herbivorous species interacts with Symbiodinium as birds, mammals, amphibians, and reptiles do with more than 700 plant species. We confirmed FLS viable cells are ubiquitous in S. viride faeces and a percentage of them survive to the grinding action of the pharyngeal jaws, and long gut pass. An average density of 33x102cells 370 mg-1 dry weight faeces is released. From these cells, the 23,12%±1,50 can maintain viable after the digestion process, which represent a significative difference between the viable and non-viable cells dispersed. Faeces size had no effect on the number of viable FLS quantified. Based on S. viride defecation rates, it is possible to infer a capacity to disperse as many as 76.000 FLS viable cells individual-1 daily on reefs. This viable cells input repopulates FLS environmental reservoirs as sediments and generates positive ecosystem feedbacks regarding reef resilience. Thus, FLS genetic connectivity maintenance and genotypic diversity increase. Also, new FLS are constantly becoming available for aposymbiotic cnidarian uptake, as those experimenting post-bleaching or other diseases in the current environmental global context. In conclusion, our results increase the understanding of S. viride functional role as a new coral reef resilience driver. S. viride conservation efforts focus on overfishing

16 regulation, and involvement of locals to participate in environmental actions are a priority for the maintenance of its ecological functions and reefs` resilience feedbacks.

Acknowledgments: We would like to thank Universidad de los Andes (Faculty of Science) and Consejo Profesional de Biología for funding this project. Also, we acknowledge Corporación para el Desarrollo Sostenible del Archipiélago de San Andrés, Providencia y Santa Catalina (CORALINA), and Buconos Diving hotel team for the assistance during field surveys. Natalia Bolaños from the Laboratorio de Ciencias Básicas Médicas at the Universidad de los Andes supported the FCM analysis process. Humberto Ibarra and his team from the Centro de Microscopía at the Universidad de los Andes contributed with microscopy methods. Finally, we kindly thank reviewers Dr. Elvira Alvarado Chacón and Dr. Emilio Realpe; besides, the Laboratorio de Biología Molecular Marina team (BIOMMAR) at the Universidad de los Andes for their valuable and constant assistance. This thesis is dedicated to parrotfish and Symbiodinium lovers wide world.

Bibliography: Adam, T. C., Burkepile, D. E., Ruttenberg, B. I., & Paddack, M. J. (2015). Herbivory and the resilience of Caribbean coral reefs: knowledge gaps and implications for management. Marine Ecology Progress Series, 520, 1–20. http://doi.org/10.3354/meps11170

Aronson, R. B., & Precht, W. F. (2006). Conservation, precaution, and Caribbean reefs. Coral Reefs, 25(3), 441–450. http://doi.org/10.1007/s00338-006-0122-9

Aswani, S., Mumby, P. J., Baker, A. C., Patrick, C., Laurence, J. M., S., S. R., & Richmond Robert H. (2015). Scientific frontiers in the management of coral reefs. Frontiers in Marine Science, 2(50), 1–13. http://doi.org/10.3389/fmars.2015.00050

Augustine, L., & Muller-Parker, G. (1998). Selective predation by the mosshead sculpin Clinocottus globiceps on the sea anemone Anthopleura elegantissima and its two algal symbionts. Limnology and Oceanography, 43(4), 711–715. http://doi.org/10.4319/lo.1998.43.4.0711

Bachman, S., & Muller-Parker, G. (2007). Viable algae released by the seastar Dermasterias imbricata feeding on the symbiotic sea anemone Anthopleura elegantissima. Marine Biology, 150, 369–375. http://doi.org/10.1007/s00227-006-0344-y

Baine, M., Howard, M., Kerr, S., Edgar, G., & Toral, V. (2007). Coastal and marine resource management in the Galapagos Islands and the Archipelago of San Andrés : Issues, problems and opportunities. Ocean & Coastal Management, 50, 148–173. http://doi.org/10.1016/j.ocecoaman.2006.04.001

Bello, C., Galetti, M., Montan, D., Pizo, M. A., Mariguela, T. C., Culot, L., … Jordano, P. (2017). Atlantic frugivory: a plant–frugivore interaction data set for the Atlantic Forest. Ecology, 98(6), 1729. http://doi.org/10.1002/ecy.1818

Bellwood, D. R., Hoey, A. S., & Hughes, T. P. (2012). Human activity selectively impacts the ecosystem roles of parrotfishes on coral reefs. Proceedings of the Royal Society B: Biological Sciences, 279(1733), 1621–1629. http://doi.org/10.1098/rspb.2011.1906 Bellwood, D. R., Hughes, T. P., Folke, C., & Nystrom, M. (2004). Confronting the coral reef crisis.

17 Nature, 429(6994), 827–833.

Bennett, S., Wernberg, T., Harvey, E. S., Santana-Garcon, J., & Saunders, B. J. (2015). Tropical herbivores provide resilience to a climate-mediated phase shift on temperate reefs. Ecology Letters, 18(7), 714–723. http://doi.org/10.1111/ele.12450

Blanco, G., Bravo, C., Pacifico, E. C., Chamorro, D., Speziale, K. L., Lambertucci, S. A., … Tella, J. L. (2016). Internal seed dispersal by parrots: an overview of a neglected mutualism. PeerJ, 4, e1688. http://doi.org/10.7717/peerj.1688

Blank, R. J. (1987). Cell architecture of the dinoflagellate Symbiodinium sp. inhabiting the Hawaiian stony coral Montipora verrucosa. Marine Biology, 94(1), 143–155. http://doi.org/10.1007/BF00392906

Bonaldo, R. M., Hoey, A. S., & Bellwood, D. R. (2014). The ecosystem roles of parrotfishes on tropical reefs. Oceanography and Marine Biology: An Anual Review, 52, 81–132.

Bruggemann, J. H., Van Kessel, A. M., Van Rooij, J. M., & Breeman, A. M. (1996). Bioerosion and sediment ingestion by the caribbean parrotfish Scarus vetula and Sparisoma viride: Implications of fish size, feeding mode and habitat use. Marine Ecology Progress Series, 134(1–3), 59–71. http://doi.org/10.3354/meps134059

Burghardt, I., & Wägele, H. (2014). The symbiosis between the “solar-powered” nudibranch Melibe engeli Risbec, 1937 (Dendronotoidea) and Symbiodinium sp. (Dinophyceae). Journal of Molluscan Studies, 80(5), 508–517. http://doi.org/10.1093/mollus/eyu043

Burkepile, D. E., Allgeier, J. E., Shantz, A. A., Pritchard, C. E., Lemoine, N. P., Bhatti, L. H., & Layman, C. A. (2013). Nutrient supply from fishes facilitates macroalgae and suppresses corals in a Caribbean coral reef ecosystem. Scientific Reports, 3(1), 1493. http://doi.org/10.1038/srep01493

Burtscher, M. M., May, L. A., Downs, C. A., & Bartlett, T. (2015). Zooxanthellae Viability Assay. In C. M. Woodley, C. A. Downs, A. W. Bruckner, J. W. Porter, & S. B. Galloway (Eds.), Diseases of Coral. John Wiley & Sons, Inc, Hoboken, NJ. http://doi.org/10.1002/9781118828502.ch39

Buysschaert, B., Byloos, B., Leys, N., Van Houdt, R., & Boon, N. (2016). Reevaluating multicolor flow cytometry to assess microbial viability. Applied Microbiology and Biotechnology, 100(21), 9037–9051. http://doi.org/10.1007/s00253-016-7837-5

Castaño-Isaza, J., Newball, R., Roach, B., & Lau, W. W. Y. (2015). Valuing beaches to develop payment for ecosystem services schemes in Colombia Seaflower marine protected area. Ecosystem Services, 11, 22–31.

Campam.gcfi.org. (2017). Welcome - Caribbean Marine Protected Areas Management. [online] Available at: http://campam.gcfi.org/CaribbeanMPA/CaribbeanMPA.php [Sept. 13 2017].

Castro-Sanguino, C. (2010). Ichthyochorous dispersal of Symbiodinium spp.: an ecological process for maintaining viable environmental pools of symbionts? Universidad de los Andes.

Castro-sanguino, C., & Sánchez, J. (2012). Dispersal of Symbiodinium by the stoplight parrotfish Sparisoma viride. Biology Letters Marine Biology, 8, 282–286.

Comeros-Raynal, M. T., Choat, J. H., Polidoro, B. A., Clements, K. D., Abesamis, R., Craig, M. T., …

18 Carpenter, K. E. (2012). The likelihood of extinction of iconic and dominant herbivores and detritivores of coral reefs: The parrotfishes and surgeonfishes. PLoS ONE, 7(7), 1–13. http://doi.org/10.1371/journal.pone.0039825

Conde, J. E., Diaz, H., & Sambrano, A. (1991). Disintegration of holothurian fecal pellets in beds of the seagrass Thalassia testudinum. Journal of Coastal Research, 7(3), 853–862.

Cunning, R., Yost, D. M., Guarinello, M. L., Putnam, H. M., & Gates, R. D. (2015). Variability of Symbiodinium communities in waters, sediments, and corals of thermally distinct reef pools in American Samoa. PLoS ONE, 10(12), 1–17. http://doi.org/10.1371/journal.pone.0145099

De Vega, C., Arista, M., Ortiz, P. L., Herrera, C. M., & Talavera, S. (2011). Endozoochory by beetles: A novel seed dispersal mechanism. Annals of Botany, 107(4), 629–637. http://doi.org/10.1093/aob/mcr013

Delgado, A. (2013). Aproximación a la viabilidad de zooxantelas del género Symbiodinium spp. presentes en las heces del pez loro Sparisoma viride. Universidad de los Andes.

Edwards, C. B., Friedlander, A. M., Green, A. G., Hardt, M. J., Sala, E., Sweatman, H. P., … Smith, J. E. (2014). Global assessment of the status of coral reef herbivorous fishes : evidence for fishing effects Proceedings of the Royal Society B: Biological Sciences, 281(20131835), 7–11. http://doi.org/http://dx.doi.org/10.1098/rspb.2013.1835

FAO. (2014). The State of World Fisheries and Aquaculture 2014.

Fitt, W. K., & Trench, R. K. (1983). The relation of diel patterns of cell division to diel patterns of motility in the symbiotic dinoflagellate Symbiodinium microadriaticum Freudenthal in culture. New Phytologyst, 94, 421–432. http://doi.org/10.1111/j.1469-8137.1983.tb03456.x

Francini-filho, R. B., Moura, R. L., Ferreira, C. M., & Coni, E. O. C. (2008). Live coral predation by parrotfishes (Perciformes: Scaridae ) in the Abrolhos Bank, eastern Brazil, with comments on the classification of species into functional groups. Neotropical Ichtyology, 6(2), 191–200.

Goatley, C. H. R., & Bellwood, D. R. (2010). Biologically mediated sediment fluxes on coral reefs: sediment removal and off-reef transportation by the surgeonfish Ctenochaetus striatus. Marine Ecology Progress Series, 415(January 2014), 237–245. http://doi.org/10.3354/meps08761

Granados-Cifuentes, C., Neigel, J., Leberg, P., & Rodriguez-Lanetty, M. (2015). Genetic diversity of free- living Symbiodinium in the Caribbean: the importance of habitats and seasons. Coral Reefs, 34(3), 927– 939. http://doi.org/10.1007/s00338-015-1291-1

Guy-Haim, T., Hyams-Kaphzan, O., Yeruham, E., Almogi-Labin, A., & Carlton, J. T. (2017). A novel marine bioinvasion vector: Ichthyochory, live passage through fish. Limnology and Oceanography Letters, 2(3), 81–90. http://doi.org/10.1002/lol2.10039

Hopkinson, C. S. (1985). Shallow-water benthic and pelagic metabolism: Marine Biology, 87(1), 19–32. http://doi.org/10.1007/BF00397002 Restrepo, J. (2007). Cultivo de las zooxantelas extraídas de las heces del pez loro, Sparisoma viride. Universidad de los Andes.

Kempf, S. C. (1991). A “primitive” symbiosis between the aeolid nudibranch Berghia verrucicornis (A. costa, 1867) and a zooxanthella. Journal of Molluscan Studies, 57, 75–85.

19 http://doi.org/10.1093/mollus/57.Supplement_Part_4.75

Krediet, C. J., Denofrio, J. C., Caruso, C., Burriesci, M. S., Cella, K., & Pringle, J. R. (2015). Rapid, Precise, and Accurate Counts of Symbiodinium Cells Using the Guava Flow Cytometer, and a Comparison to Other Methods. PLoS ONE, 10(8), 1–19. http://doi.org/10.1371/journal.pone.0135725

Lee, C. S., Sheng, Y., Yeo, W., & Sin, T. M. (2012). Bleaching Response of Symbiodinium (Zooxanthellae): Determination by Flow Cytometry. Cytometry Part A, 81A, 888–895. http://doi.org/10.1002/cyto.a.22111

Levin, R. A., Suggett, D. J., Nitschke, M. R., van Oppen, M. J. H., & Steinberg, P. D. (2017). Expanding the Symbiodinium (Dinophyceae, Suessiales) Toolkit Through Protoplast Technology. Journal of Eukaryotic Microbiology, 64(5), 588–597. http://doi.org/10.1111/jeu.12393

Littman, R. A., van Oppen, M. J. H., & Willis, B. L. (2008). Methods for sampling free-living Symbiodinium (zooxanthellae) and their distribution and abundance at Lizard Island (Great Barrier Reef). Journal of Experimental Marine Biology and Ecology, 364(1), 48–53. http://doi.org/https://doi.org/10.1016/j.jembe.2008.06.034

Lobel, P. S. (1981). Trophic biology of herbivorous reef fishes: alimentary pH and digestive capabilities. Journal of Fish Biology, 19(4), 365–397. http://doi.org/10.1111/j.1095-8649.1981.tb05842.x

MacNeil, M. A., Graham, N. A. J., Cinner, J. E., Wilson, S. K., Williams, I. D., Maina, J., … McClanahan, T. R. (2015). Recovery potential of the world’s coral reef fishes. Nature, 520(7547), 341– 344. http://doi.org/10.1038/nature14358

Marin, A., & Ros, J. (1991). Presence of intracellular zooxanthellae in mediterranean nudibranchs. Journal of Molluscan Studies, 57, 87–101. http://doi.org/10.1093/mollus/57.Supplement_Part_4.87

Maruyama, T., & Heslinga, G. A. (1991). Fecal discharge of zooxanthellae in the giant clam Tridacna derasa, with reference to their in situ growth rate. Marine Biology, 127, 473–477. http://doi.org/10.1007/s002270050035

McAfee, S. T., & Morgan, S. G. (1996). Resource use by five sympatric parrotfishes in the San Blas Archipelago, Panama. Marine Biology, 125(3), 427–437. http://doi.org/10.1007/BF00353255

McCarthy, D. A., & Macey, M. G. (2007). Flow cytometry: principles and applications. (M. G. Macey, Ed.). Totowa, N.J. : Humana Press,. Retrieved from http://www.springerlink.com/openurl.asp?genre=book&isbn=978-1-58829-691-7

Mora, C. (2008). A clear human footprint in the coral reefs of the Caribbean. Proceedings. Biological Sciences / The Royal Society, 275(1636), 767–773. http://doi.org/10.1098/rspb.2007.1472

Mora, C. (2015). Ecology of Fishes on Coral Reefs. Cambridge: Cambridge University Press. http://doi.org/DOI: 10.1017/CBO9781316105412

Muller-Parker, G. (1984). Dispersal of Zooxanthellae on Coral Reefs by Predators on Cnidarians. Biological Bulletin, 167(1), 159–167.

Muller-Parker, G., D’Elia, C. F., & Cook, C. B. (2015). Interactions Between Corals and Their Symbiotic Algae. In C. Birkeland (Ed.), Coral Reefs in the Anthropocene (pp. 99–116). Dordrecht: Springer

20 Netherlands. http://doi.org/10.1007/978-94-017-7249-5_5

Mumby, P. J., Dahlgren, C. P., Harborne, A. R., Kappel, C. V, Micheli, F., Brumbaugh, D. R., … Gill, A. B. (2006). Fishing, trophic cascades, and the process of grazing on coral reefs. Science (New York, N.Y.), 311(5757), 98–101. http://doi.org/10.1126/science.1121129

Neo, M. L., Eckman, W., Vicentuan, K., Teo, S. L. M., & Todd, P. A. (2015). The ecological significance of giant clams in coral reef ecosystems. Biological Conservation, 181, 111–123. http://doi.org/10.1016/j.biocon.2014.11.004

Nitschke, M. (2015). The free-living Symbiodinium reservoir and scleractinian coral symbiont acquisition. The University of Queesland.

Nitschke, M. R., Davy, S. K., & Ward, S. (2016). Horizontal transmission of Symbiodinium cells between adult and juvenile corals is aided by benthic sediment. Coral Reefs, 35(1), 335–344. http://doi.org/10.1007/s00338-015-1349-0

Pakeman, R. J., Digneffe, G., & Small, J. L. (2002). Ecological correlates of endozoochory by herbivores. Functional Ecology, 16(3), 296–304. http://doi.org/10.1046/j.1365-2435.2002.00625.x

Pakeman, R. J., Engelen, J., & Attwood, J. P. (1999). Rabbit endozoochroy and seedbank build-up in an acidic grassland. Plant Ecology, 145(1), 83–90. http://doi.org/10.1023/A:1009864030814

Pandolfi, J. M. (2015). Deep and complex ways to survive bleaching, 8–9.

Papandreou, S. (2013). Flow cytometry : principles, methodology and applications. Hauppauge, New York : Nova Science Publishers, Inc.,.

Papoutsoglou, E. S., & Lyndon, A. R. (2006). Digestive enzymes along the alimentary tract of the parrotfish Sparisoma cretense. Journal of Fish Biology, 69(1), 130–140. http://doi.org/10.1111/j.1095- 8649.2006.01082.x

Perry, C. T., Kench, P. S., O’Leary, M. J., Morgan, K. M., & Januchowski-Hartley, F. (2015). Linking reef ecology to island building: Parrotfish identified as major producers of island-building sediment in the Maldives. Geology, 43(6), 503–506. http://doi.org/10.1130/G36623.1

Plass-Johnson, J. G., Ferse, S. C. A., Jompa, J., Wild, C., & Teichberg, M. (2015). Fish herbivory as key ecological function in a heavily degraded coral reef system. Limnology and Oceanography, 60(4), 1382– 1391. http://doi.org/10.1002/lno.10105

Pollux, B. J. A. (2011). The experimental study of seed dispersal by fish (ichthyochory). Freshwater Biology, 56(2), 197–212. http://doi.org/10.1111/j.1365-2427.2010.02493.x

Poore, A. G. B., Campbell, A. H., Coleman, R. a., Edgar, G. J., Jormalainen, V., Reynolds, P. L., … Emmett Duffy, J. (2012). Global patterns in the impact of marine herbivores on benthic primary producers. Ecology Letters, 15(8), 912–922. http://doi.org/10.1111/j.1461-0248.2012.01804.x Porto, I., Granados, C., Restrepo, J. C., & Sánchez, J. A. (2008). Macroalgal-associated dinoflagellates belonging to the genus Symbiodinium in caribbean reefs. PLoS ONE, 3(5), 1–5. http://doi.org/10.1371/journal.pone.0002160

Poulicek, M., Roberty, S., Plaza, S., & Ladriere, O. (2010). Symbiodinium sp . can stay alive through the

21 gut and in the faeces of cnidaria predators: the case of Coralliophilla meyendorffi and Anemonia viridis. In Rapp. Comm. int. Mer Médit. (p. 637).

Pozdnyakov, I., & Skarlato, S. (2012). Dinoflagellate amphiesma at different stages of the life cycle. Protistology, 7(2), 108–115.

Price, S. A., Wainwright, P. C., Bellwood, D. R., Kazancioglu, E., Collar, D. C., & Near, T. J. (2010). Functional innovations and morphological diversification in parrotfish. Evolution, 64(10), 3057–3068. http://doi.org/10.1111/j.1558-5646.2010.01036.x

Puyana, M., Acosta, A., Bernal-Sotelo, K., Velásquez-Rodríguez, T., & Freddy, R. (2015). Spatial scale of cyanobacterial blooms in Old Providence Island, Colombian Caribbean. Universitas Scientiarum, 20(1), 83–105. http://doi.org/10.11144/Javeriana.SC20-1.sscb

Reyes-Nivia, M. C., Garzón-Ferreira, J., & Rodríguez-Ramírez, A. (2004). Depredación de coral vivo por peces en el Parque Nacional Natural Tayrona, Caribe colombiano. Revista de Biología Tropical, 52(4), 883–895. Retrieved from http://www.scielo.sa.cr/scielo.php?script=sci_abstract&pid=S0034- 77442004000400007&lng=en&nrm=iso&tlng=en

Rotjan, R. D., & Lewis, S. M. (2005). Selective predation by parrotfishes on the reef coral Porites astreoides. Marine Ecology Progress Series, 305(September 2017), 193–201. http://doi.org/10.3354/meps305193

Sánchez, J. A., Gil, M. F., Chasqui, L. H., & Alvarado, E. M. (2004). Grazing dynamics on a Caribbean reef-building coral. Coral Reefs, 23(4), 578–583. http://doi.org/10.1007/s00338-004-0419-5

Sazima, C., Bonaldo, R. M., Krajewski, J. P., & Sazima, I. (2005). The Noronha wrasse: a “jack-of-all- trades” follower. Aqua, Journal of Ichthyology and Aquatic Biology, 9(3), 97–108. http://doi.org/10.1007/s10641-006-9123-3

Seavy, B. E., & Muller-Parker, G. (2002). Chemosensory and feeding responses of the nudibranch Aeolidia papillosa to the symbiotic sea anemone Anthopleura elegantissima. Invertebrate Biology, 121(2), 115–125. http://doi.org/10.1111/j.1744-7410.2002.tb00052.x

Shikang, S., Fuqin, W., & Yuehua, W. (2015). Does the passage of seeds through frugivore gut affect their storage: A case study on the endangered plant Euryodendron excelsum. Scientific Reports, 5(1), 11615. http://doi.org/10.1038/srep11615

Stambler, N. (2011). Zooxanthellae: The Yellow Symbionts Inside Animals BT. In Z. Dubinsky & N. Stambler (Eds.), Coral Reefs: An Ecosystem in Transition (pp. 87–106). Dordrecht: Springer Netherlands. http://doi.org/10.1007/978-94-007-0114-4_7

Staniforth, R. J., & Cavers, P. B. (1977). The Importance of Cottontail Rabbits in the Dispersal of Polygonum Spp. Journal of Applied Ecology, 14(1), 261–268. http://doi.org/10.2307/2401841

Strychar, K. B., Coates, M., Sammarco, P. W., Piva, T. J., & Scott, P. T. (2005). Loss of Symbiodinium from bleached soft corals Sarcophyton ehrenbergi, Sinularia sp. and Xenia sp. Journal of Experimental Marine Biology and Ecology, 320(2), 159–177. http://doi.org/10.1016/j.jembe.2004.12.039

Suárez-Esteban, A., Delibes, M., & Fedriani, J. M. (2013). Barriers or corridors? The overlooked role of unpaved roads in endozoochorous seed dispersal. Journal of Applied Ecology, 50(3), 767–774.

22 http://doi.org/10.1111/1365-2664.12080

Takabayashi, M., Adams, L. M., Pochon, X., & Gates, R. D. (2012). Genetic diversity of free-living Symbiodinium in surface water and sediment of Hawai‘i and Florida. Coral Reefs, 31(1), 157–167. http://doi.org/10.1007/s00338-011-0832-5

Thompson, K., Band, S. R., & Hodgson, J. G. (1993). Seed Size and Shape Predict Persistence in Soil. Functional Ecology, 7(2), 236–241. http://doi.org/10.2307/2389893

Thornhill, D. J., Howells, E. J., Wham, D. C., Steury, T. D., & Santos, S. R. (2017). Population genetics of reef coral endosymbionts (Symbiodinium, Dinophyceae). Molecular Ecology, 26(10), 2640–2659. http://doi.org/10.1111/mec.14055

Tol, S. J., Jarvis, J. C., York, P. H., Grech, A., Congdon, B. C., & Coles, R. G. (2017). Long distance biotic dispersal of tropical seagrass seeds by marine mega-herbivores. Scientific Reports, 7(1), 4458. http://doi.org/10.1038/s41598-017-04421-1

Tolimieri, N. (1998). The relationship among microhabitat characteristics recrultme t and adult abundance in the stoplight parrotfish, Sparisoma viride, at three spatial scales. Bulletin of Marine, 62(1), 253–268.

Venera-Ponton, D. E., Diaz-Pulido, G., Rodriguez-Lanetty, M., & Hoegh-Guldberg, O. (2010). Presence of Symbiodinium spp. in macroalgal microhabitats from the southern Great Barrier Reef. Coral Reefs, 29(4), 1049–1060. http://doi.org/10.1007/s00338-010-0666-6

Vermeij, M. J. A., van der Heijden, R. A., Olthuis, J. G., Marhaver, K. L., Smith, J. E., & Visser, P. M. (2013). Survival and dispersal of turf algae and macroalgae consumed by herbivorous coral reef fishes. Oecologia, 171(2), 417–425. http://doi.org/10.1007/s00442-012-2436-3

Wilkinson, D. M., Lovas-Kiss, A., Callaghan, D. A., & Green, A. J. (2017). Endozoochory of large bryophyte fragments by waterbirds. Cryptogamie, Bryologie, 38(2), 223–228. http://doi.org/10.7872/cryb/v38.iss2.2017.223