# Agglomerated loricae of the tintinnids Codonella, Codonellopsis and Dictyocysta from North Atlantic, tropical Pacific and Southern Ocean waters

Gustaaf M Hallegraeff, Ruth S Eriksen, Karine Leblanc

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Gustaaf M Hallegraeff, Ruth S Eriksen, Karine Leblanc. Agglomerated loricae of the tintinnids Codonella, Codonellopsis and Dictyocysta from North Atlantic, tropical Pacific and Southern Ocean waters. Journal of Plankton Research, 2025, 47 (5), pp.fbaf030. 10.1093/plankt/fbaf030. hal-05208862

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Microzooplankton

# Agglomerated loricae of the tintinnids *Codonella*, *Codonellopsis* and *Dictyocysta* from North Atlantic, tropical Pacific and Southern Ocean waters

Gustaaf M. Hallegraeff<fnref n="1" />,*, Ruth S. Eriksen<fnref n="1" />,<fnref n="2" />,<fnref n="3" /> and Karine Leblanc<fnref n="4" />

## ABSTRACT

Some 30 species of planktonic tintinnids agglomerate coccoliths and diatom fragments on to their loricae, which have fascinated scientists for centuries. We here provide semi-quantitative scanning electron microscopic observations on tintinnid lorica agglomerations by the small Southern Ocean *Codonellopsis* cf. *soyai* Hada, intermediate sized North Atlantic *Codonellopsis pusilla* (Cleve) Kofoid and Campbell, and larger Southern Ocean *Dictyocysta lepida* Ehrenberg and tropical Pacific *Codonella galea* Haeckel. Smaller tintinnids appeared to non-selectively use ubiquitous *Emiliania huxleyi* liths, but when absent could shift to ambient diatom fragments, while larger tintinnids preferentially used heavier coccoliths of *Calcidiscus*, *Coccolithus* and *Helicosphaera* adding up to an estimated 25% extra lorica weight. Lorica agglomerations appeared more prominent in colder waters compared to warmer tropical waters. Selective feeding in tintinnids is closely linked to their swimming behavior, and possible benefits from agglomerated loricae for ballasting and slowing of swimming speed are discussed.

**KEYWORDS:** tintinnids; coccoliths; *Codonellopsis* cf. *soyai*; *Codonellopsis pusilla*; *Dictyocysta lepida*; *Codonella galea*

## INTRODUCTION

Tintinnids are ubiquitous planktonic ciliates that live inside a protective solid shell, called a lorica, with diagnostic morphology. These microzooplankton create a feeding current that draws food particles toward their mouth using a crown of continuously beating oral cilia (Wandel and Holzman, 2022). Their proteineous loricae have fascinated scientists for centuries (Haeckel, 1873). Cementing of mineral particles for test formation is well documented also for foraminifera, phaeodarians and thecate amoebae (Stout and Walker, 1976; Gowing, 1989; Bowser and Bernhard, 1993), but tintinnids stand out for their capacity to also use coccoliths and diatom fragments. Tintinnid loricae can be either hyaline, or agglutinated with biogenic (coccoliths, diatom frustules) or non-biogenic particles (sand grains, mineral flakes). If the agglutinated particles are comprised exclusively of mineral grains the lorica is called arenaceous (“sandy”), but if decorated with particles of mineral and/or biological origin the lorica is called agglomerated (Gold and Morales, 1976). Adhering particles can cover the entire surface or only the bowl, but mostly leave the collar uncovered. Whether agglomerated particles are rejected food particles or directly

captured on to sticky loricae is not clear. Details of the process of lorica formation and agglutination also remain incompletely understood. Agglutination appears to stop after construction of the bowl when the lorica hardens and loses its stickiness (Agatha et al., 2013). That is, biogenic particles are captured during only a very brief time slot immediately after tintinnid cell division. In the classical taxonomic monographs on tintinnids by Kofoid and Campbell (1929, 1939) these authors illustrated by line drawings 753 tintinnid species from the eastern tropical Pacific, of which 22 species (3%) carried coccoliths and only a single species (*Laackmanniella naviculaefera*) carried diatoms. The increasing use of scanning electron microscopy (SEM) has added many more agglomerated tintinnid species to this list, but more importantly this allowed for resolution of the precise identity of agglomerating biogenic materials (Burns, 1983; Takahashi and Ling, 1984; Winter et al., 1986; Wasik et al., 1996; Henjes and Assmy, 2008).

Intriguingly, only selected tintinnid genera such as *Acanthostomella*, *Codonella*, *Codonellopsis*, *Dictyocysta*, *Laackmanniella*, *Stenosemella* and *Tintinnopsis* appear to be able to agglutinate biogenic particles. Furthermore, not all species within a genus

<footnote n="1">¹Institute for Marine and Antarctic Studies, University of Tasmania, 20 Castray Esplanade, Battery Point, Hobart, TAS 7004, Australia</footnote>

<footnote n="2">²CSIRO Environment, 3-4 Castray Esplanade, Battery Point, TAS 7004, Australia</footnote>

<footnote n="3">³Australian National Algae Culture Collection, CSIRO National Collections and Marine Infrastructure, 3-4 Castray Esplanade, Battery Point, TAS 7004, Australia</footnote>

<footnote n="4">⁴Aix Marseille Université, Université de Toulon, CNRS, IRD, MIO UM 110, 163 avenue de Luminy, 13288 Marseille, France</footnote>

<footnote>*Corresponding author: gustaaf.hallegraeff@utas.edu.au</footnote>

<footnote>Corresponding editor: John Dolan</footnote>
agglutinate (Kofoid and Campbell, 1929, 1939). Much discussion has centered on whether the adhered particles, either food remnants or suspended particles from the water column, reflect the nature of the ecosystem in which the lorica was built. At present, only three semi-quantitative studies exist on tintinnid lorica agglomerations, two of which focused on the small tintinnid *Stenosemella* using coccoliths and/or diatoms (Winter et al., 1986; Henjes and Assmy, 2008), and one study that focused on diatom agglomerations (Wasik et al., 1996). In the present work, we add new semi-quantitative SEM observations from the North Atlantic, tropical Pacific and Southern Ocean Oceans: (i) to expand the list of tintinnid taxa with agglomerated loricae; (ii) identify the nature of coccolith and diatom biogenic particles on tintinnids in different ocean regions; as a first step (iii) to better understand the possible ecological advantage of lorica agglomeration.

# MATERIAL AND METHODS

## Sample collection and preparation

Water samples from the Southern Ocean Time Series (SOTS) voyage were collected on board RV Investigator in May 2023 using a CTD rosette sampler at depths between 10 and 80 m, filtered directly on board ship on 0.8 µm Nuclepore filters, rinsed with deionized water, airdried, coated with platinum (5 nm layer thickness), and examined with a Hitachi SU70 field emission scanning electron microscope (FESEM). Samples collected using the RV Sprightly cruises SP16/80 and SP 3A/82 in the Coral Sea in November 1980 and March 1982, respectively, employed a 37 µm freefall plankton net from surface to 200 m depth. Unpreserved samples were collected on to polycarbonate filters on board ship, airdried and examined with a JEOL-35C Scanning Electron Microscope. The MobyDick cruise took place in February–March 2018 around Kerguelen Island, in the Indian sector of the Southern Ocean. Phytoplankton net and bottle net samples (Agusti et al., 2015) were collected at four locations during the late summer bloom. Light microscopy images were captured on board on live samples less than 30 min after sampling, while SEM pictures were acquired 3 months later on uncoated polycarbonate filters with a Phenom SEM. Samples from the APERO campaign off the Porcupine Abyssal Plain in the North Atlantic were collected in June–July 2023 using Niskin bottles, phytonets (35 µm mesh) (from 0 to 200 m), particle traps (from 50 to 1000 m) and deep nets (20 µm mesh) from 200 to 4000 m. For each sample, two aliquots were fixed with acid Lugol and basic formalin for observation by light microscopy (Nikon TE-200) while a third sample was filtered directly on board ship on a polycarbonate filter, then coated with gold (8 nm thickness) for observation by Phenom ProX SEM. Samples on the Tonga cruise were collected in October–December 2019 from Niskin, phytonets (35 µm mesh) and deep nets (20 µm) (from 200 to 2000 m), and observed on a Nikon TE-200 light microscope and a Phenom Pro SEM. The scientific cruise details of the sample collections used in this study have been summarized in Table I and the geographic locations are mapped in Fig. 1.

<figure><img src="image_2.png" /><figcaption>Fig. 1. Sample locations for SEM observations of agglomerated tintinnid loricae used in the present study: 1. North Atlantic (Porcupine Abyssal Plain); 2. Kerguelen Island Plateau; 3. SOTS station; 4. Tonga-Kermadec Arc. Supplementary material was obtained from 5. The Coral Sea; and 6. Indian Ocean. The locations of previous semi-quantitative tintinnid lorica studies by 7. Winter et al. (1986) and 8. Henjes and Assmy (2008) are also indicated. Red dots (locations 1–4) represent the main sample materials used, black numbers (locations 5–6) supplementary material, and white dots (7–8) location of literature data.</figcaption></figure>

## Tintinnid lorica assessment

Scanning micrographs of loricae (10 to 35 per species) were printed full-size on A4 paper and individual coccoliths or diatoms contoured, counted and measured. While scanning micrographs have the advantage of resolution of the identity of biogenic materials, when loricae are too densely covered with agglutinated particles this can prevent identification of the tintinnid species. We therefore only selected loricae for which tintinnid identity was unambiguous. Similarly, SEMs typically only allow for inspection of half of the lorica surface, and we had to assume (but see Fig. 26) that both lateral sides were identical.

In order to estimate the extra lorica ballast from carrying coccoliths, we used the following conversion factors. The ellipsoidal liths of *Emiliania huxleyi* (3–4 µm length) and *Gephyrocapsa muelleri* (3.5–5 µm length) were nearly identical in size, *Syracosphaera* ellipsoidal liths ranged up to 5.6 µm long, *Coccolithus* liths were up to 7.8 µm long, the discoidal liths of *Calcidiscus* were 9 µm in diameter, and *Helicosphaera* liths were 6–12 µm long. We used coccolith weights estimated by Beaufort (2005) and Young and Ziveri (2000) to calculate the lith weights as follows: *Emiliania* 4 pg CaCO₃ per lith, *Gladitholithus* (4 pg), *Syracosphaera* (10 pg), *Umbilicosphaera* (16 pg), *Florisphaera* (20 pg), *Rhabdosphaera* (60 pg), *Helicosphaera* (140 pg), *Calcidiscus* (124 pg) and *Coccolithus* (152 pg). Different morphotypes of *E. huxleyi* were not discriminated in these calculations. To estimate the weight of diatom fragments, we estimated their biovolumes (e.g. *Pseudo-nitzschia* fragment 25 µm³, *Chaetoceros* setae 40 µm³, *Fragilariopsis* fragment 63 µm³, *Thalassiothrix* fragment 100 µm³, *Eucampia* cribrum fragment 250 µm³, *Coscinodiscus* cribrum fragment 375 µm³), and converted biovolumes to silica using the equations of Conley
<table><caption>**Table I:** Sample locations for SEM observations of agglomerated tintinnid loricae used in the present study. Locations are mapped in Fig. 1.</caption><thead><tr><th>Locality</th><th>Coordinates</th><th>Dates</th><th>Sampling device</th><th>Sample preparation</th><th>Tintinnid taxa</th></tr></thead><tbody><tr><td colspan="6"><strong>Tropics</strong></td></tr><tr><td>Indian Ocean<br>(Fig. 1, site 6)</td><td>along 110°E,<br>11.5–23°S</td><td>IIOE-2 RV Investigator<br>IN2019 voyage V03<br>May–June 2019</td><td>20 µm net, 0–20 m<br>depth</td><td>Lugol and buffered<br>formalin preserved,<br>Hitachi FESEM</td><td>Dictyocysta lepida</td></tr><tr><td>Coral Sea<br>(Fig. 1, site 5)</td><td>SP 16/80; SP3A/82;<br>149–152°E,<br>15–21°S</td><td>RV Sprightly cruises<br>SP16/80; SP3A/80<br>outer edge Great Barrier<br>Reef; November 1980;<br>March 1982</td><td>37 µm freefall net,<br>0–200 m depth</td><td>Unpreserved,<br>Nuclepore filters,<br>Hitachi FESEM</td><td>Codonella galea,<br>Dictyocysta<br>lepida,<br>Dictyocysta<br>spinosa</td></tr><tr><td>Tropical Pacific<br>(Fig. 1, site 4)</td><td>Tonga-Kermadec Arc;<br>165°E–165°W;<br>19–22°S</td><td>TONGA expedition<br>October–December<br>2019</td><td>Niskin bottles,<br>phytonets (35 µm<br>mesh), deep nets<br>(20 µm) (from 200<br>to 2000 m)</td><td>Lugol preserved.<br>Phenom ProX SEM</td><td>Codonella galea</td></tr><tr><td colspan="6"><strong>Temperate waters</strong></td></tr><tr><td>North Atlantic<br>(Fig. 1, site 1)</td><td>Porcupine Abyssal<br>Plain; 48°50′N,<br>016°30′W</td><td>APERO expedition,<br>June–July 2023</td><td>Niskin bottles,<br>phytonets (35 µm<br>mesh), 0–200 m,<br>particle traps,<br>50–1000 m, deep<br>nets (20 µm mesh),<br>200–4000 m</td><td>Unpreserved,<br>polycarbonate filter,<br>Phenom ProX SEM<br>on untreated and<br>uncoated filters</td><td>Dictyocysta<br>lepida,<br>Codonellopsis<br>pusilla</td></tr><tr><td colspan="6"><strong>Southern Ocean</strong></td></tr><tr><td>(Fig. 1, site 2)</td><td>Kerguelen Island<br>plateau; 67–75°E,<br>49–52°S</td><td>MOBYDICK cruise<br>February–March 2018</td><td>35 µm mesh<br>phytoplankton net<br>and bottle net<br>(holding 20 µm<br>mesh) samples</td><td>Unpreserved,<br>polycarbonate filter<br>Phenom ProX SEM<br>on untreated and<br>uncoated filters</td><td>Codonellopsis cf.<br>soyai</td></tr><tr><td>(Fig. 1, site 3)</td><td>Southern Ocean Time<br>Series<br>(42.84–47.48°S;<br>139.8–147.6°E)</td><td>RV Investigator<br>IN2023-voyage V03,<br>May 2023</td><td>CTD rosette<br> sampler, 10–80 m<br>depth</td><td>Unpreserved,<br>Nuclepore filters,<br>Hitachi FESEM</td><td>Codonellopsis<br>pusilla<br>Dictyocysta lepida</td></tr></tbody></table>

*et al.* (1989); roughly 0.0005 pmole silica per µm³, with 1 pmole equaling to 28 pg silica. For small intact diatoms such as *Fragilariopsis pseudonana* we used a biovolume of 100 µm³ corresponding to 1.4 pg silica per cell.

## Water column coccolithophorids

For the APERO North Atlantic samples, semi-quantitative coccolith counts were conducted on ~0.25 mL subsamples from 1 L diluted plankton net collections, counting 1/4 of a 25 mm filter by Phenom SEM. Southern Ocean (SOTS) ambient coccolithophorid counts derive from deep (1000–3800 m) sediment trap samples preserved in mercury chloride, of which subsamples were mounted on microscope slides, and coccospheres and coccoliths counted using a Nikon Eclipse 80i polarized light microscope at 1000× magnification (Rigual Hernández *et al.*, 2020.) Unfortunately, no ambient coccolithophorid data were available for the Indian Ocean, Tonga and MobyDick (Kerguelen Plateau) cruises.

# RESULTS

## Codonellopsis cf. soyai

A small tintinnid closely resembling *Codonellopsis soyai* Hada described from the Indian sector of the Southern Ocean (Hada, 1970, Fig. 48; type illustration reproduced as present Fig. 2) was common off the Kerguelen Island Plateau in February–March 2018 (Fig. 1, location 2). Diagnostic features were the flask-shaped lorica composed of an ovoid bowl [33–44 µm (rarely 49–52 µm) long, 32–40 µm wide] and transparent (Fig. 3) subcylindrical collar (7.5–8.5 µm high), composed of 3–5 rings, with numerous (7–12) ovate fenestrae present mostly in the middle two rings (Figs 5–8). The surface of the obovoid bowl, 21–24 µm oral diameter, as observed on broken or corroded loricae, was smooth, but always densely covered with coccolithophorids and/or diatoms or diatom fragments. Both longer (Fig. 3) and shorter loricae (Fig. 4) co-occurred, with comparable oral diameter, but these all exhibited identical agglutination patterns.

Of 11 loricae examined, six were predominantly covered by *E. huxleyi* (Fig. 5; based on genetic arguments also referred to as
<figure><img src="image_3.png" /><figcaption>**Figs 2–10.** Fig. 2: Type illustration of *Codonellopsis soyai* from the Indian Ocean sector of the Southern Ocean (reproduced from Hada, 1970, Fig. 48); Figs 3–10. Tintinnid loricae of C. cf. *soyai* from the Kerguelen Island Plateau. Figs 3–4. Light micrographs of long and short loricae with agglutinated *Fragilariopsis* diatom fragments and whole cells; Figs 5–10. Scanning Electron Micrographs of loricae ornamented by (Fig. 5) predominantly *Emiliania huxleyi* (Eh) coccoliths or (Figs 6–8) broken diatom fragments of Cos = *Coscinodiscus/Asteromphalus*, Eu = *Eucampia*, Fra = *Fragilariopsis kerguelensis*, Thx = *Thalassiothrix antarctica*, PsN=Pseudo-nitzschia, Chae = *Chaetoceros* setae. Figs 9 and 10. High magnification of diatom fragment agglutination. Scale bars: 25 µm (Figs 3–4); 10 µm (Figs 6–8).</figcaption></figure>
<table><caption>**Table II:** Biogenic particles on loricae of Codonellopsis soyai from the Kerguelen Island Plateau. Emiliania lith cover (pg CaCO₃/lorica) and diatom fragments (as pg silica/lorica).</caption><thead><tr><th rowspan="2">Emiliania<br>pg CaCO<sub>3</sub>/lorica<br>(% of total ballast)<br>Kerguelen Island<br>Plateau</th><th colspan="6">Diatom fragments pg silica/lorica</th><th rowspan="2">Total ballast<br>pg/lorica</th></tr><tr><th>Fragila<br>riopsis</th><th>Pseudo-<br>nitzschia</th><th>Chaeto<br>ceros<br>setae</th><th>Thalassio<br>thrix</th><th>Coscino<br>discus<br>cribra</th><th>Eucam<br>pia<br>cribra</th></tr></thead><tbody><tr><td>1488 (100)</td><td></td><td></td><td></td><td></td><td></td><td></td><td>1488</td></tr><tr><td>880 (99)</td><td>5</td><td>0.7</td><td></td><td>5.6</td><td></td><td></td><td>891</td></tr><tr><td>960 (92)</td><td></td><td></td><td></td><td></td><td>84</td><td></td><td>1044</td></tr><tr><td>464 (98)</td><td>2</td><td>1</td><td></td><td>8</td><td></td><td></td><td>475</td></tr><tr><td>528(96)</td><td></td><td></td><td>13</td><td></td><td>11</td><td></td><td>552</td></tr><tr><td>288(97)</td><td>5</td><td>1</td><td>2</td><td></td><td></td><td></td><td>296</td></tr><tr><td>116(64)</td><td>2</td><td></td><td></td><td>25</td><td>32</td><td>7</td><td>180</td></tr><tr><td>88(56)</td><td></td><td></td><td></td><td>11</td><td>52</td><td>7</td><td>158</td></tr><tr><td>88(79)</td><td></td><td></td><td>13</td><td></td><td>10</td><td></td><td>111</td></tr><tr><td>0 (0)</td><td></td><td></td><td>15</td><td></td><td>10</td><td></td><td>25</td></tr><tr><td>0 (0)</td><td>26</td><td>2</td><td>7</td><td></td><td></td><td></td><td>35</td></tr><tr><td>445 ± 461<br>(73 ± 36%) (n = 11)</td><td>5 of 11</td><td>4 of 11</td><td>5 of 11</td><td>4 of 11</td><td>6 of 11</td><td>2 of 11</td><td>477 ± 454</td></tr></tbody></table>

Gephyrocapsa huxleyi, Bendif et al., 2023; Wheeler et al., 2023) but five others were almost exclusively covered with diatom fragments (Figs 6–10), including broken Chaetoceros setae, fragments of Fragilariopsis, Pseudo-nitzschia, Thalassiothrix, and fragments of Coscinodiscus/Asteromphalus and heavily silicified frustules of Eucampia antarctica. The diatom fragments were agglutinated directly onto the loricae and difficult to quantify. Rarely, complete cells of the diatom Fragilariopsis were found attached (Fig. 3). For loricae dominated by Emiliania, the coccolithophorid load was estimated to be up to 1488 pg per lorica (Table II). For loricae covered exclusively by diatom fragments this was estimated to be much lower as 35–296 pg per lorica.

## Codonellopsis pusilla

Tintinnids corresponding to Codonellopsis pusilla (Cleve) Kofoid and Campbell (type illustration by Cleve, 1899, p.970 as Codonella pusilla reproduced as present Fig. 11) were abundant in samples from the North Atlantic Porcupine Abyssal Plain (Fig. 1, location 1). This species had a clavate to ovate lorica (Figs 11–17), 29–35 µm wide, 32–35 µm bowl length, with a transparent collar of 4–5 rings, oral diameter 18–23 µm, and with 3–5 large elliptical fenestrae (Figs 14–17) in a single middle ring of the collar. The bowl was widest near its middle, with a hemispherical or sometimes pointed aboral end. Non-agglomerated corroded loricae exhibited a distinct hexagonal surface pattern on the lorica wall (Fig. 14) also described for the related Codonellopsis contracta Kofoid and Campbell (type illustration reproduced in Fig. 12). Non-agglutinated loricae were clavate in shape but heavily agglomerated loricae more ovoid or even bulbous in outline. Of 35 loricae examined from the North Atlantic, the majority were covered by different morphotypes of E. huxleyi coccoliths (Fig. 15), with small amounts of Syracosphaera pulchra liths regularly present. Only five loricae were dominated by Calcidiscus (positioned in a distinct band; Fig. 16), three were dominated by Coccolithus

braarudii/pelagicus (Fig. 13) with the diatoms F. pseudonana (4–5 µm long) on top of the bowl, with a single cell of Shionodiscus oestrupii or Nitzschia bicapitata (14 µm long; not shown) also observed embedded on top (Fig. 17). A typical lorica carried an estimated average coccolith weight of 1689 ± 542 pg, but the capture by 30% of the North Atlantic tintinnid population of Calcidiscus, Helicosphaera and Coccolithus liths (Fig. 18) raised this value by 3 to 6 fold up to 9696 pg (mean 2585 ± sd 2390 pg). The addition of small diatoms on the top of the bowl at most added only some 3% to lorica weight (Table III). Corroded loricae showed that larger Coccolithus and Calcidiscus liths did not create a canopy over smaller liths. By contrast, small Fragilariopsis diatoms were superimposed upon Emiliania liths. North Atlantic lorica agglomerated with Calcidiscus and Coccolithus liths were estimated to be heavier than those that carried accessory diatoms (Table III). Comparable loricae were also observed at the SOTS station (Fig. 1, location 3; Fig. 19, n = 13), oral diameter 18–19 µm, length of bowl 35–36 µm, but these were either completely covered by Calcidiscus, or with smaller Emiliania and sometimes Alisphaera mainly covering the 20–30% of the bottom of the bowl (Fig. 20). A single whole diatom cell of N. bicapitata was observed on only 1 out of 13 loricae studied. The Southern Ocean loricae, heavily ballasted by Calcidiscus, were estimated to weigh an extra 10 540 ± 3537 pg (Table III).

## Codonella galea

Loricae identified as Codonella galea Haeckel had a large pot-shaped body, oral diameter 40–72 µm (average ± sd of 51 ± 10 µm), 60–113 µm (77 ± 16 µm) long, divided into a funnel-shaped collar with finely denticulate margin and a subspherical bowl, separated by a constricted neck (Figs 21–24). Codonella differs from Codonellopsis in that the collar is not hyaline (Fig. 22) but mineral coated and includes agglutinated biogenic particles. It differs from Codonaria in the absence of a suboral cone. A single lorica from the Coral Sea (Fig. 1,
<figure><img src="image_4.png" /><figcaption>**Figs 11–18.** Fig. 11: Type illustration of *Codonellopsis pusilla* from Newfoundland Banks (Cleve, 1899, p.970, Fig. 3). Fig. 12: Type illustration of *Codonellopsis contracta* from the Peruvian Current showing sparse fenestrae (Fn) (Kooid and Campbell, 1939, Fig. 147); Figs 13–18: Tintinnid loricae of *C. pusilla* from the North Atlantic (Porcupine Abyssal Plain); Scanning electron micrographs showing variations in bowl shape according to heavy coccolith agglutination (Figs 13 or 14). Empty corroded clavate bowl with hexagonal surface structure from an unpreserved sample filtered directly on board ship. Lorice ornamented by (Fig. 15) predominantly *Emiliania huxleyi* with *Syracosphaera pulchra* (Sy) and *Gephyrocapsa muelleri* (Gm); (Fig. 16) *Calcidiscus leptoporus* (Ca); and (Fig. 17) agglutinated diatoms *Shinodiscus oestrupii* (So), *Thalassiosira* (Ths) and complete diatom cells of *Fragilariopsis pseudonana* (Fpn). Fig. 18 High magnification of large *Coccolithus* (Co) and *Helicosphaera carteri* liths, compared to small *E. huxleyi* liths (Eh). Scale bars: 20 µm (Figs14–17); 30 µm (Fig. 13).</figcaption></figure>
<figure><img src="image_5.png" /><figcaption>**Figs 19–24.** Figs 19–20: Scanning Electron Micrographs of *Codonellopsis pusilla* from the Southern Ocean, agglomerated with predominantly *Calcidiscus leptoporus* (Ca) liths, and (Fig. 20) *E. huxleyi* and *Alisphaera unicornis* (Al) liths only at the bottom of the bowl. Figs 21–24: *Codonella galea* from the Coral Sea (Fig. 21) and Tonga waters. (Figs 22–24). Fig. 21. Scanning micrograph of lorica from Coral Sea agglomerated mainly with *Florisphaera* (Fl) and sparse *Emiliania* (eh) and *Umbellosphaera irregularis* (Ubi) liths only around the collar. Fig. 22. Light micrograph of *C. galea* showing agglutinations also on the non-transparent collar. Figs 23–24. Scanning Electron Micrographs of *C. galea* from Tonga, adorned with *Umbellosphaera tenuis* (Ubt), *Syracosphaera mediterranea* (Sm), with (Fig. 24) *Gladiolithus* (Gl) around the collar large *Rhabdosphaera* (Rh) liths concentrated mostly toward the bottom of the bowl. Scale bars: 10 µm (Fig. 19); 20 µm (Figs 23 and 24); 25 µm (Fig. 21).</figcaption></figure>
<figure><img src="image_6.png" /><figcaption>**Figs 25–30.** Figs 25–27: Scanning electron micrographs of *Dictyocysta lepida* from the SOTS station. Fig. 25. Bowl heavily agglutinated with *Calcidiscus* (Ca) with *Helicosphaera* (He) only at the top of the bowl. Fig. 26. Bottom view of lorica showing symmetrical coverage of liths. Fig. 27. High magnification of *Calcidiscus* lith deeply embedded on to the lorica. Fig. 28: Lorica of *D. lepida* from Tonga waters agglomerated with mostly *U. tenuis* (Ubt) and *Rhabdosphaera* (Rh) liths. Fig. 29: Lorica from the Indian Ocean with *Helicosphaera*, *Gephyrocapsa* (Ge) and *Umbellosphaera* in both proximal (top of bowl) and distal positions (bottom of bowl). Fig. 30: Detail of lorica of the related *Dictyocysta spinosa* from the Coral Sea with deeply embedded *Gephyrocapsa oceanica* liths onto the bowl. Scale bars: 10 µm (Fig. 26); 15 µm (Figs 25, 28, 29).</figcaption></figure>
<table><caption>**Table III:** Biogenic particles agglomerated on the loricae of Codonellopsis pusilla from the North Atlantic and Southern Ocean. Estimated as pg CaCO₃/lorica for individual coccoliths and as total pg liths.</caption><thead><tr><th>Emiliania</th><th>Gephyrocapsa<br>oceanica</th><th>Calcidiscus</th><th>Coronosphaera</th><th>Umbellosphaera</th><th>Helicosphaera</th><th>Coccolithus</th><th>Total pg<br>CaCO<sub>3</sub>/lorica</th><th>Diatom<br>Fragments</th></tr></thead><tbody><tr><td>Calcidiscus dominated-type (n = 10; Fig. 16)</td><td></td><td>35 ± 11</td><td>14 ± 7</td><td>0</td><td>0</td><td>0</td><td>5008 ±<br>1305</td><td>Fragilariopsis pseudonana<br>4 of 11 loricae</td></tr><tr><td>North Atlantic</td><td>104 ± 33</td><td>31 ± 22</td><td></td><td></td><td></td><td></td><td></td><td></td></tr><tr><td>Coccolithus dominated-type (n = 5; Fig. 13)</td><td></td><td></td><td></td><td></td><td></td><td></td><td></td><td></td></tr><tr><td>81 ± 38</td><td></td><td></td><td>0</td><td>0</td><td>3 ± 4</td><td>40 ± 12</td><td>6966 ±<br>1990</td><td>Nitzschia bicapitata<br>2 of 5 loricae</td></tr><tr><td>E. pseudonana diatom-type (n = 9; Fig. 17)</td><td></td><td></td><td></td><td></td><td></td><td></td><td></td><td></td></tr><tr><td>187 ± 344</td><td></td><td>1 ±<br>31</td><td>8 ± 310</td><td>0</td><td>0</td><td>0</td><td>1228 ±<br>3518</td><td>34 ± 35 E. pseudonana pg<br>silica per lorica</td></tr><tr><td>19 ±<br>312</td><td></td><td></td><td></td><td></td><td></td><td></td><td></td><td></td></tr><tr><td>Others (n = 11; Fig. 15)</td><td></td><td></td><td></td><td></td><td></td><td></td><td></td><td></td></tr><tr><td>213 ± 53</td><td></td><td>13 ± 13</td><td>16 ± 10</td><td>0</td><td>0</td><td>0</td><td>1276 ±<br>323</td><td></td></tr><tr><td>Total (n = 35)</td><td>170 ± 70</td><td>17 ± 16</td><td>7 ± 14</td><td>11 ± 10</td><td>1 of 35</td><td>1 of 35</td><td>2585 ±<br>35 of 35</td><td>11 of<br>35 loricae</td></tr><tr><td>Southern Ocean</td><td>50 ± 40</td><td>1 of 13</td><td>83 ± 30</td><td>1 of 13</td><td>0</td><td>0</td><td>10 540 ±<br>3537</td><td>N. bicapitata<br>1 of 13 loricae</td></tr><tr><td>Time Series<br>(n = 13)</td><td></td><td></td><td></td><td></td><td></td><td></td><td></td><td></td></tr></tbody></table>

location 5) was densely agglomerated with small liths resembling *Florisphaera profunda* (Fig. 21; compare Young et al., 2003, plate 39, Fig. 3) with *Umbellosphaera irregularis* and *Emiliania* liths only present around the top of the collar. Liths resembling *Gladiolithus flabellatus* were found around the collar of Tonga loricae (Fig. 24; compare Young et al., 2003, plate 39, Fig. 11). A distinctive feature of this tintinnid was the use of prolate *Rhabdosphaera* rhabdoliths (up to 12 µm long) on 6 out of 12 loricae from tropical Tonga (Fig. 1, location 4), mostly on the bottom of the bowl (Fig. 24), accompanied by *Umbellosphaera*, *Syracosphaera mediterranea* and sparse *Emiliania*. Coccolith loadings for this large tintinnid were low and variable in the range $1312 \pm 706$ pg (Table IV).

## Dictyocysta lepida

Dictyocysta species represent among the most elaborate tintinnid lorica architectures known, composed of a goblet-shaped bowl, with a tall cylindrical collar, 35–41 µm oral diameter, consisting of beams surrounding 6–10 large windows in one or several rows (Figs 25–29). We here focus on the ubiquitous *Dictyocysta lepida* Ehrenberg (*Dictyocysta elegans* Ehrenberg subsp. *lepida* sensu Balech, 1959) with the collar made up of a single row of windows (Figs 25, 28, 29). The rounded bowl carried 6–7 large subcircu- lar fenestrae covered by a membrane, but sometimes (3 out of 18 loricae) completely obscured by agglomerating coccoliths. With other individuals, coccoliths could be seen deeply embedded in the framework comprising the wall of the bowl (Fig. 27). *D. lep- ida* loricae from the SOTS station in May 2023 (Fig. 1, location 3) carried predominantly (80–100% of area) *Calcidiscus* liths, with sparse *Helicosphaera* also present (Fig. 25). Total coccolith weight for this cold water population was 19 922 ± 5 287 pg (n = 18). Loricae from tropical Tonga waters (Fig. 1, location 4) were more diversely ornamented with mostly *Umbellosphaera* and *Rhabdosphaera* (Fig. 28), or *E. huxleyi* (75%) with *Umbel- losphaera* and sparse *Calciosolenia* (not shown). Loricae from the Indian Ocean (Fig. 1, location 6; Fig. 29) carried *Umbellosphaera*, *Helicosphaera* and *Gephyrocapsa oceanica*. The related species *Dictyocysta spinosa* Kofoid and Campbell from the Coral Sea (Fig. 1, location 5) exhibited exclusively *G. oceanica* coccoliths deeply embedded on the bottom of the bowl (Fig. 30). Tropical loricae carried significantly less coccolith weight $2428 \pm 2141$ pg per cell (n = 8), approximately eight times less than their cold- water counterparts (Table V).

Figure 31 summarizes the identity and abundance of agglomer- erating coccoliths and diatoms for the five tintinnid species investi- gated in this study, in which we included comparable quan- titative observations previously published for *Stenosemella* by Winter et al. (1986) and Henjes and Assmy (2008). Tintinnid taxa studied are arranged according to increasing lorica diameter (as an important correlate for prey size and feeding rates; Dolan, 2010), and coccoliths are similarly arranged according to increas- ing lith size.

## DISCUSSION

### Taxonomy of *C. pusilla*, *C. soyai* and *C. galea*

The small tintinnid *C. pusilla* (Cleve) Kofoid and Campbell, 1929 has been widely reported from diverse localities such as
<table><caption>Table IV: Biogenic particles agglomerated on the loricae of Codonella galea. Estimated as pg CaCO₃/lorica for individual coccoliths and as total liths.</caption><thead><tr><th></th><th>Emiliania huxleyi</th><th>Calcidiscus</th><th>Calciosolenia</th><th>Florisphaera</th><th>Gladiolithus</th><th>Umbilicosphaera</th><th>Umbellosphaera</th><th>Syracosphaera pulchra/ mediterranea</th><th>Rhabdosphaera</th><th>Total pg CaCO3/lorica</th></tr></thead><tbody><tr><td rowspan="7">Tonga</td><td>88</td><td>0</td><td>0</td><td>280</td><td>0</td><td>184</td><td>0</td><td>20</td><td>360</td><td>932</td></tr><tr><td>172</td><td>0</td><td>0</td><td>0</td><td>0</td><td>504</td><td>0</td><td>90</td><td>120</td><td>1058</td></tr><tr><td>0</td><td>0</td><td>0</td><td>0</td><td>0</td><td>121</td><td>12</td><td>190</td><td>390</td><td>704</td></tr><tr><td>2000</td><td>868</td><td>0</td><td>0</td><td>0</td><td>0</td><td>0</td><td>0</td><td>0</td><td>2869</td></tr><tr><td>0</td><td>0</td><td>0</td><td>0</td><td>0</td><td>640</td><td>0</td><td>50</td><td>1500</td><td>2190</td></tr><tr><td>96</td><td>0</td><td>0</td><td>200</td><td>0</td><td>416</td><td>144</td><td>60</td><td>300</td><td>1216</td></tr><tr><td>0</td><td>0</td><td>16</td><td>0</td><td>0</td><td>680</td><td>0</td><td>10</td><td>660</td><td>1366</td></tr><tr><td rowspan="3">Total (n=9)</td><td>96</td><td>0</td><td>0</td><td>0</td><td>0</td><td>256</td><td>0</td><td>90</td><td>0</td><td>594</td></tr><tr><td>96</td><td>0</td><td>0</td><td>0</td><td>0</td><td>264</td><td>0</td><td>135</td><td>390</td><td>885</td></tr><tr><td>366± 672</td><td>1 of 9</td><td>1 of 9</td><td>2 of 9</td><td>0</td><td>330± 233</td><td>2 of 9</td><td>80± 56</td><td>413± 432</td><td>1312± 706</td></tr><tr><td>Coral Sea (n=1)</td><td>88</td><td>0</td><td>0</td><td>400</td><td>880</td><td>88</td><td>22</td><td>0</td><td>0</td><td>1472</td></tr></tbody></table>

<figure><img src="image_7.png" /><figcaption>Fig. 31. Summary diagram of tintinnid agglomerations with coccoliths and diatom or diatom fragments examined in the present study, supplemented with observations on *Stenosemella* by Rassoulzadegan (1980), Winter *et al.* (1986), Gowing and Garrison (1992) and Henjes and Assmy (2008). The five tintinnid taxa studied are arranged according to increasing lorica diameter (left) for the specimens observed, and agglomerating coccoliths are similarly arranged according to increasing lith size (top). Large blue circles mark tintinnid preferences for coccolith species, red circles designate the ability to use diatom whole cells or diatom fragments, while the yellow circles indicate the use of large prolate rhabdoliths exclusively by the two largest tintinnids; + indicate present.</figcaption></figure>
<table><caption>Table V: Biogenic particles (% of total liths) agglomerated on the loricae of Dictyocysta lepida. Estimated as pg CaCO₃/lorica for individual coccoliths and as total liths.</caption><thead><tr><th></th><th>Emiliania huxleyi</th><th>Gephyrocapsa</th><th>Calcidiscus leptoporus</th><th>Umbilicosphaera foliosa</th><th>Umbellosphaera tenuis</th><th>Helicosphaera carteri</th><th>Calciosolenia</th><th>Rhabdosphaera</th><th>Total pg CaCO₃/lorica</th></tr></thead><tbody><tr><td>Southern Ocean Time Station (n = 18)</td><td>1 of 18</td><td>0</td><td>155 ± 46</td><td>0</td><td>0</td><td>10 of 18</td><td>0</td><td>0</td><td>19 922 ± 5 287</td></tr><tr><td>Indian Ocean (n = 5)</td><td>46 ± 63</td><td>2 of 5</td><td>0</td><td>2 of 5</td><td>23 ± 15</td><td>17 ± 17</td><td>0</td><td>0</td><td>3 248 ± 2 299</td></tr><tr><td>Tonga (n = 3)</td><td>74 ± 69</td><td>0</td><td>0</td><td>0</td><td>35 ± 17</td><td>0</td><td>1 of 3</td><td>1 of 3</td><td>2 428 ± 2 141</td></tr></tbody></table>

the Subarctic (Cleve, 1899), Antarctic (Sassi and Melo, 1986; Petz, 2005) and even the Mediterranean (Balech, 1959, his Figs 81–84). This points to either an extremely eurythermal species but more likely taxonomic confusion. *Codonellopsis pusilla* was originally described from Newfoundland Banks (at water temperatures of 9–14°C) and placed in *Codonella* (Cleve, 1899) but later formally transferred to *Codonellopsis* by Kofoid and Campbell (1929, p.87) because of its a high annular collar distinctively set off from the bowl. This transfer is often credited to Jörgensen (1924) who described the new genus *Codonellopsis* (but without designating a type species), and while he mentioned in the text C. pusilla var.ovata (p.98) and *Codonella pusilla* (p.101), this taxon is not discussed, no basionym is provided, and no formal transfer instigated. This species was described as having an obovoid “clavate” bowl, 14–15 µm oral diameter, 35–48 µm long, with a cylindrical, annulated collar with “a few transverse rings”. Fenestrae were not illustrated (present type illustration Fig. 11) nor mentioned by Cleve (1899) but could have been easily overlooked. Critically, the surface of the lorica was described as “with coarse rounded pores, some of which are hexagonally framed”. This distinctive feature fits our material from the North Atlantic (Fig. 14). Balech (1959) in his treatise of Mediterranean tintinnids referred to C. pusilla as a highly variable species and pointed to similarities with C. *contracta*. Kofoid and Campbell, which was described as having a wall with “secondary structure (Kofoid and Campbell, 1929, Fig. 147) and with “one or more elliptical fenestrae in the lower part of the spiral” (present Fig. 12). The key difference between these two taxa is the more inflated bowl and hemispherical aboral end of C. *contracta*. It has not been widely recognized however that dense biogenic agglomerations can significantly obscure and change the shape and width of the bowl (compare Figs 14 and 16). Balech (1959) also pointed out that the warm-water Mediterranean forms he identified as C. pusilla could well belong to the poorly defined taxon *Codonellopsis monacensis* (Rampi) Balech, originally described from Mediterranean waters as *Stenosemella monacense* (Rampi, 1950, p.4, Fig. 1).

By contrast, our material from the Kerguelen Island Plateau did not match C. pusilla but most closely fits C. soyai Hada described from the Indian sector of the Southern Ocean (Hada, 1970). Diagnostic features were the flask-shaped lorica with obovoid bowl and subcylindrical collar. Critically the surface of the obovoid bowl was described as smooth (not with a hexagonal pattern as in C. pusilla), The collar was flared at the oral end, composed of 3–5 spiral turns “with many small ovate fenestellae”. As in C. pusilla. The surface of the bowl was described as having “some coccoliths as adhering foreign particles” but Hada (1970, Fig. 48) illustrated in great detail a striking pattern that retrospectively could be interpreted as agglomerated diatom or diatom fragments (compare present Figs 2 and 3). In our Antarctic material we observed both longer (49–52 µm, Fig. 3) and shorter loricae (33–44 µm, Fig. 4), but with identical oral diameter, transdiameter and agglutination patterns. We did not attempt to separate these morphotypes. The lorica lengths of our longer morphotype matched that reported by Hada, 1970 (50–52 µm). Petz (2005) reported both C. soyai and C. pusilla from the Antarctic, but also suggested C. soyai to be probably conspecific with C. pusilla with which we do not
agree. In addition to striking differences in number of fenestrae in the collar rings, the distinctive agglomerations of coccoliths and diatom fragments documented here between these two taxa also support their discrimination. The variability of lorica dimensions and shapes of smaller tintinnids remain poorly defined (summarized in Supplementary Table 1). At present no molecular sequences are available for these or related taxa (Bachy, 2012), which is critical to resolve the biogeography of these small tintinnids.

The tropical tintinnid we here refer to as *C. galea* agrees in all respect with the emended description by Kofoid and Campbell (1939) of *C. galea* of 78–120 µm lorica length (emended from Kofoid and Campbell, 1929), except for the incidence of smaller cells of 60–113 µm length (77 average ± 16 µm standard deviation). We note that other authors similarly included smaller forms under this name such as 54–120 µm long (Marshall, 1969) and 73–100 µm long (Komarovsky, 1959, from the Red Sea). Critically, Kofoid and Campbell, 1939 described by LM comparable coccolith agglutination as we observed from *Umbilicosphaera, Syracosphaera mediterranea, Rhabdosphaera clavigera* (except for not being able to resolve the smaller *Florisphaera* and *Gladiolithus*-like liths, which we detected by SEM).

**Do tintinnids have a preference for particular coccoliths?**

In the present work, we investigated tintinnid populations for which we had suitable material available in terms of numbers of individuals, preservation and unambiguous tintinnid species identifications to semi-quantitatively assess lorica variations at a single location. We here address tintinnids that primarily utilize coccoliths rather than diatoms, which we will expand on in a separate communication. Wasik et al. (1996) previously documented abundance and species compositions of diatoms adhering to the loricae of Antarctic *Laackmaniella naviculaefera, Codonellopsis gaussi, C.balechi* and Baltic Sea *Tintinnopsis lobiancoi*. These authors concluded that certain diatoms (*Fragilariopsis, Thalassiosira*) were actively selected and it is not a rule that tintinnids only use those which are dominant. Agglomerations of morphologically variable diatoms and diatom fragments are more challenging to quantify than coccolith agglomerations, also calling for attention to the attachment of living diatoms in a possible symbiotic association (e.g. Armbrecht et al., 2017; Gómez, 2020). The summary diagram in Fig. 31 demonstrates how the two smallest, cold-water tintinnids *Stenosemella* and *Codonellopsis* cf. *soyai* (17–23 µm oral diameter) appeared to non-selectively use the ubiquitous smallest coccolithophorids *Emiliania huxleyi* or *Gephyrocapsa muelleri* (3–4 µm lith length), most often attached to the lorica with the proximal shield. They rarely attached sparse larger liths such as *Calcidiscus* or *Coccolithus* (the latter illustrated by Winter et al., 1986 for *Stenosemella* from the Agulhas Current). However, when coccolithophorids were sparse or absent, these tintinnids shifted to using almost exclusively ubiquitous diatom fragments. This shift was first demonstrated by Henjes and Assmy (2008) for *Stenosemella* from the Antarctic Polar Front, and here was also suggested for *C. cf. soyai* from the Kerguelen Plateau. Admittedly, our material did not cover time series observations nor data on ambient plankton communities. Tintinnids are thought not to be able to mechanically crush diatoms, hence in all likelihood

these microzooplankton capitalize on the leftovers from active feeding on diatom blooms by larger copepods or krill. The ubiquitous nanoplankton *Emiliania* is unique in being able to produce multiple layers of coccoliths and prolifically shed liths into ambient seawater, a feature not known however from any other coccolithophorid species. This raises the question of how tintinnids are able to dismantle large coccolithophorid cells because agglomerated liths are almost always intact. The intermediate sized tintinnid *C. pusilla* from the North Atlantic and the Southern Ocean, oral diameter 20–23 µm but up to 35 µm bowl length, was able to use small *Emiliania, Gephyrocapsa* and *Syracosphaera* liths but if available appeared to preferentially select larger and heavier *Calcidiscus* and *Coccolithus* liths (5–15 µm diameter). Ambient water column coccolithophorid assemblages in the North Atlantic (APERO cruise) were comprised in decreasing order of numerical abundance of *Emiliania, Calcidiscus, Gephyrocapsa muelleri, Syracosphaera pulchra* and *Coccolithus*. Similarly, coccolithophorid assemblages in sediment traps at the SOTS station were dominated by *Emiliania*, with *Calcidiscus* at most 20% of total coccolithophores and *Coccolithus* rare (Findlay and Giraudeau, 2000; Rigual Hernández et al., 2020). Diatom fragments were rarely used by this tintinnid species but more common was the agglutination on the top of the bowl of small intact diatom cells of *Fragilariopsis pseudonana* (4–5 µm) or *Nitzschia bicapitata* (14 µm long). The latter has also been reported from, e.g. *Acanthostomella minutissima* from New Zealand (Burns, 1983). This strategy constitutes a transition to tintinnids such as *C. gaussi, Laackmanniella* and *Tintinnopsis* that exclusively use diatoms (Wasik et al., 1996) many of them alive (Armbrecht et al., 2017). Of interest is that the freshwater *Codonella cratera* Leidy, thriving in lake environments without coccolithophorids, uses small discoid diatoms such as *Cyclotella ocellata* (5 µm diameter) of a size and shape comparable to *Emiliania* liths (Bettighofer, 2024). Similarly, small *C. pusilla* like loricae from the Mediterranean have been observed to carry mixtures of *Emiliania/Gephyrocapsa* coccoliths as well as minute (3 µm diameter) *Minidiscus* diatom cells (image available at https://ecotaxaobs-vlfr.fr/prj/15396). This strongly points to size selection of biogenic materials irrespective of their taxonomic identity.

Not surprisingly, the largest tintinnids studied here, *D. lepida* and *C. galea*, 35–65 µm oral diameter, housing the strongest swimming ciliates, exhibited the capacity to capture the greatest biodiversity of liths, again with a strong preference for the larger coccoliths of *Calcidiscus, Helicosphaera, Coccolithus, Umbellosphaera* and also the ability to capture *Rhabdosphaera* liths (up to 12 µm long). This unique latter feature was already noted by Kofoid and Campbell (1939) for *C. galea* and *Codonaria australis* (Kofoid and Campbell) Kofoid and Campbell, and confirmed by Takahashi and Ling (1984) also for *Codonellopsis minor* (Brandt) Kofoid and Campbell and by Bachy (2011) for *Ceratocephala orthoceras* (Haeckel) Jörgensen. Such prolate liths are preferentially positioned at the bottom of the bowl (Figs 24 and 28). Similarly, with *D. lepida* the same funnel-shaped *Umbellosphaera* liths could be positioned in distal view on top of the bowl, but deposited in proximal view at the bottom of the bowl (Fig. 29) with the protruding stems adding to surface roughness. Small lighter liths (e.g. *Florisphaera, Gladiolithus*) and diatoms tended
to be positioned mostly close to the mouth of the bowl (Figs 17 and 24). On some loricae, e.g. of *Codonellopsis elongata* from Palau (Young et al., 2022, JRYSEM-275-22) the coverage of spiraling rows of asymmetrical *Helicosphaera* liths strongly resembles the spiraling dynamics of ciliar beating. On other loricae heavier liths can be deposited on the middle and leaving the bottom of the bowl for smaller liths (Figs 16 and 19). These patterns suggests hydrodynamic sorting of liths jettisoned away from the cytostomal region by ciliar beating.

## Possible function of agglomerated biogenic particles

Numerous explanations have been proposed for the functions of agglomerated tintinnid loricae, including: (i) protective armor to reduce its susceptibility to copepod predation; (ii) ballast to quickly sink away from predators; (iii) enhancing capture of food particles around the oral cilia, and (iv) modification of swimming behavior by tethering to detritus (Jonsson et al., 2004) or diatom frustules (reviewed by Agatha et al., 2013). Limited data on sinking rates of tintinnids (Suzuki and Taniguchi, 1995) indicate that small hyaline loricae such as *Acanthostomella* sink at 0.25–2.08 m day⁻¹ while two species with agglutinated loricae, *Tintinnopsis ampla* Hada and *Tintinnopsis beroidea* Stein exhibited sinking rates of 1.90 and 15.9 m day⁻¹. The latter was covered with coccolithophorids, mostly *Emiliania*. Takahashi (1982) provided an estimate for the dry weight of loricae from a mixture of the medium-size *Codonellopsis minor* and larger *C. biedermanii* (Brandt) Kofoid and Campbell as 40 000 ± 20 000 pg per individual. From published images by Henjes and Assmy (2008) we estimated a coccolith load for the smallest tintinnids *Stenosemella* of 1500 to 1700 pg and for our C. *cf*. *soyai* of 880–1488 pg. By contrast, for *C. pusilla* in the Southern Ocean ballasted by heavy *Calcidiscus*, this could add an extra 10 540 ± 3537 pg (i.e. an extra 25% of lorica weight). By contrast, ballasting of the large tropical Tonga *C. galea* was insignificant at 1312 ± 3706 pg. Similarly, tropical Indian Ocean tropical *D. elegans* loricae carried significantly less coccolith weight of 2428 ± 2141 pg per cell, compared to their Southern Ocean counterparts of 19 922 ± 5287 pg. These observations suggest that tintinnid loricae agglomerations may be quantitatively more important in more viscous colder waters compared to less viscous warmer tropical waters. Agglomerations with solid calcium carbonate coccoliths consistently added more weight to tintinnid loricae than empty broken siliceous diatom frustules could achieve (Tables II–V). Loricae agglomerations however may serve purposes other than ballasting, which has also been suggested as a strategy to sink away from predators. At low Reynolds numbers seawater is viscous and the effect of lorica surface roughness on swimming speed has not yet been explored. Instead, most studies on ciliate swimming behavior have all been confined to the smooth-loricate tintinnids *Favella* and *Tintinnopsis* or loricate ciliates *Strombidium* and *Strombilidinium* (Montagnes, 2013).

Feeding and swimming in tintinnids are tightly linked by their cilia creating feeding currents that bring prey to the cytostomal region. Tintinnids can modify feeding behavior by modulating cilia kinematics (Wandel and Holzman, 2022). Probability of prey encounter increases with predator size, prey size and density. Tintinnids prefer prey of roughly 20–30% of the size of

their loricae diameter (Dolan et al., 2013) and prefer spherical instead of prolate particles. The preferential capture in our study by larger tintinnids of larger discoid *Calcidiscus*, *Coccolithus* and *Helicosphaera* liths and rarely prolate *Rhabdosphaera* liths confirm this. By contrast the smaller tintinnids appeared to use either abundant ambient small *Emiliania* liths or diatom fragments more passively (Fig. 31). When captured, prey may be rejected, or moved into the cytostomal region, where they are processed and digested, and indigestible parts egested. Whether agglomerated particles are rejected food particles or directly captured onto sticky loricae is not clear. Typical intact coccospheres of *Emiliania* are 5–12 µm in diameter (Henderiks, 2008), which is too big for ingestion by *Stenosemella*, *C. soyai* or *C. pusilla*. However, coccospHERE diameters for *Calcidiscus* are 10–13 µm, for *Coccolithus* 10–30 µm and *Helicosphaera* 9–16 µm, which is in the range of capture for the larger *D. lepida* and *C. galea*. On the other hand, the coccospHERE diameter of *Rhabdosphaera* of 20–35 µm is too much even for these larger tintinnids. Tintinnids that build agglomerated loricae offer fascinating opportunities for analysis by high-speed videography of feeding and swimming (Wandel and Holzman, 2022) and by using photogrammetry of surface roughness to estimate drag coefficients (Barton et al., 2010). The next step should be to challenge cultured or hand-picked tintinnids from freshly collected field samples with mixtures of microalgal cultures and fluorescent beads of different sizes, shapes and weights to advance our understanding of the ecological advantages of tintinnid loricae agglomeration.

## ACKNOWLEDGEMENTS

We thank all scientists, captain and crew of R/Vs *L'Atalante*, *Investigator*, *Marion-Dufresne* and *Pourquoi Pas*? for their cooperative work at sea. We thank Drs D. Vaulot and I. Probert for access to the Phenom-Pro SEM at the Roscoff Biological Station (France) for all SEM images from these three programs. Prof. Martina Doblin kindly gave us access to *Investigator* IN2019-V03 Indian Ocean samples. We thank the late Assoc. Prof. Maret Vesk from the Electron Microscope Unit of Sydney University and Dr Karsten Goemann and Dr Sandrin Feig from the Central Science Laboratories of the University of Tasmania for expert assistance with electron microscopy on the Australian samples.

## FUNDING

The MOBYDICK–THEMISTO cruise was supported by the French oceanographic fleet, the French Agence Nationale de la Recherche, AAPC-201B program, ANR-1B-CE01-001G, and Research program of INSU-CNRS LEFE/CYBER. The APERO cruise was funded by and LEFE/CYBER. The TONGA cruise GEOTRACES GPpr14, was funded by the TGIR Flotte Océanographique Française, the A-MIDEx of Aix-Marseille University, the LEFE/CYBER and GMMC program and ANR-18-CE01-0016. The Southern Ocean Time Series (SOTS) samples were collected under the auspices of the Integrated Marine Observing System (IMOS) which is enabled by the Australian National Collaborative Research Infrastructure Strategy (NCRIS). It is operated by a consortium of institutions as an unincorporated joint venture, with the University of
Tasmania as Lead Agent. This research was supported by a grant of sea time on RV Investigator from the CSIRO Marine National Facility (https://ror.org/01mae9353).

## SUPPLEMENTARY DATA

Supplementary data can be found at *Journal of Plankton Research* online.

## DATA AVAILABILITY

All SEM images used in this work have been deposited on a dedicated Ecotaxa platform focused on tintinnid loricae with agglomerated biogenic particles (https://ecotaxaobs-vlfr.fr/prj/15396). We encourage other workers to add their images of agglomerated tintinnid loricae to stimulate collaboration.

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<footnote>© The Author(s) 2025. Published by Oxford University Press. This is an Open Access article distributed under the terms of the Creative Commons Attribution License (https://creativecommons.org/licenses/by/4.0/), which permits unrestricted reuse, distribution, and reproduction in any medium, provided the original work is properly cited.</footnote>

<footnote>Journal of Plankton Research, 2025, Vol. 47, No. 5, Ba030</footnote>

<footnote>https://doi.org/10.1093/plankt/jba030</footnote>

<footnote>Original Article</footnote>