The pilot Naked Clam modular aquaculture system we developed is shown in Fig. 1, and comprises wooden panels in static (non-circulating) aerated aquaria. It was effective in allowing us to assess the relative assimilation efficacy of different feed types. This included a microencapsulated feed which consisted of an algal formulation surrounded by a waxy coating (see Methods). Faecal count analyses revealed that the mean faecal weight from Naked Clams fed on the wood + microcapsule diet (0.26 ± 0.05 (SE) mg) was significantly greater than that from Naked Clams fed on wood only (0.11 ± 0.01 (SE) mg) or wood + Shellfish Diet 1800 (0.13 ± 0.01 (SE) mg) (ANOVA, F(2,6) = 6.27, p < 0.05; Tukey’s HSD p < 0.05) (Fig. 2a). Faecal production rates were also significantly lower in Naked Clams fed on wood + microcapsules (15.0 ± 1.5 pellet individual⁻¹ day⁻¹) compared to those fed on wood + Shellfish Diet 1800 (49.0 ± 9.0 pellet individual⁻¹ day⁻¹) (ANOVA, F(2,6) = 10.71, p < 0.05; Tukey’s HSD p < 0.01) (Fig. 2b). There was however no significant difference in faecal weight per day between the diet types (ANOVA, F(2,6) = 2.02, p > 0.05) (Fig. 2c). Combined, these three results indicate that whilst there was no overall difference in assimilation rate between diet types (shown by the same faecal weight per day), the retention time for microencapsulated feed in the gut was longer, indicated by the heavier faecal pellet weight and lower production rate from Naked Clams fed on the microencapsulated feed versus those fed on the Shellfish Diet.Fig. 1

An 82 mm Teredo navalis individual, removed from the wood, is shown in (a). Most of the white meat extends beyond the valves as indicated by a yellow V in (a) – hence the name ‘Naked Clams’. Naked Clams can be grown in wood using a simple, modular aquaculture system as shown in (b). The siphons of Naked Clams are the only structures that extend beyond the wooden burrows, and are outlined by yellow dashed lines and magnified in (c). Note the fragments of waste wood dust (frass) covering the wooden panel, shown by the blue arrows, indicating active feeding and growth on wood. This, together with the ripe gonad shown by the red arrow in (a), demonstrates the efficacy of our modular aquaculture system in rearing healthy animals.

The Scanning Electron Microscopy - Energy Dispersive Spectroscopy (SEM-EDS) data shows that Naked Clams supplemented with microcapsules display clear differences in faecal material (frass), both compositionally and elementally, compared with control animals fed on a diet solely composed of wood (Fig. 3). SEM images show animals feeding only on wood produced uniform shredded wood fragments in their waste (Fig. 3a), compared with frass that was larger and clumpier in animals enriched with a microcapsule feed (Fig. 3b). This corroborates the findings of our feed assimilation efficacy trials, where animals with a microencapsulated supplemented diet produced heavier faecal pellets (Fig. 2a) and no noticeable pseudofaeces. Subsequent EDS analysis of faecal material from these microcapsule supplemented animals displayed approximately 5× the amount of calcium with minor amounts of silica and sulphur also detected (Fig. 3c–f).Fig. 3

SEM images of Naked Clam faeces at 250× magnification (scale bar = 50 µm) for animal feeding on wood only (a) and wood plus microcapsules (b). Elemental mapping overlay image of faeces for animal feeding on wood only (c) and wood plus microcapsules (d), with raw SEM image displayed in the top right corner. EDS elemental spectra from (c) and (d), shown in (e) and (f) respectively.

We were successful in performing the first formal nutritional profile of a Naked Clam (T. navalis). The fatty acid profile, carbohydrate, and protein content, as well as the organic micronutrient vitamin B₁₂ (hereafter B₁₂), was compared across our three feeding regimes, and also against that of the commercially farmed blue mussel (Mytlius edulis, referred to as blue mussels hereon). This analysis was also the first to determine the B₁₂ content of Naked Clams, and a second trial of a lipid analysis methodology developed specifically for bivalve tissue (de-coagulation of bivalve tissues prior to lipid extraction).

Basic biochemical composition analyses revealed that Naked Clams had a higher ash content than blue mussels, and hence lower levels of the metabolite groups protein, fat, and carbohydrate (Fig. 4). Proteins were the dominant metabolite group in both Naked Clams and mussels. Importantly, supplementation with the microencapsulated feed led to a significantly greater protein content than feeding on wood alone, with values of 29% and 22.5% per unit dry weight (DW) (ANOVA, F(1,22) = 14.04, p < 0.05; Tukey–Kramer’s HSD, p < 0.05). Carbohydrate content ranged between 16–19% w/w, while lipid content was 3.5–3.8% w/w.Fig. 4

a Protein, carbohydrate, lipid, and (b) vitamin B₁₂ content (expressed as a fraction of total dry weight) of Naked Clams fed with wood only (brown), Naked Clams fed with wood + microcapsules (red), and commercially farmed blue mussels (Mytilus edulis) (blue). Data are expressed as mean +/− standard deviation (SD) of technical replicates (n = 12 for Naked Clams, n = 4 for blue mussels). Letters x, y, and z indicate post-hoc test outcomes, where shared letters indicate no significant difference between groups.

Analyses revealed that Naked Clams had significantly higher levels of B₁₂ than blue mussels (Fig. 4). The B₁₂ content of Naked Clams fed on wood + microcapsules was 142 ± 9 (SE) µg B₁₂ per 100 g DW, significantly greater than that of blue mussels at 81 ± 9 (SE) µg per 100 g DW (ANOVA, F(1,13) = 9.87, p < 0.01; Tukey–Kramer’s HSD, p < 0.01). There was however no significant difference in B₁₂ levels between Naked Clams fed on wood only or wood + microcapsules; supplementation with microencapsulated feed did not positively influence the B₁₂ content.

Fatty acid composition assessments revealed that Naked Clams contained a greater content of long chain saturated fatty acids than blue mussels, and that supplementation with microencapsulated feed increases the content of polyunsaturated fatty acids (PUFAs) EPA (Eicosapentaenoic acid, 20:5(n-3)) and DHA (Docosahexaenoic acid, 22:6(n-3)) (Fig. 5). Mass spectrometry revealed that in Naked Clams reared on either wood only or wood + microcapsules, the saturated FAMEs stearic acid (C18:0) and palmitic acid (C16:0) were dominant, followed by the monounsaturated FAMEs oleic acid (C18:1) and palmitoleic acid (C16:1) in high abundance (Fig. 5a). Supplementation of the wood only diet with microencapsulated feed appeared to have an impact on the polyunsaturated acid distribution and content (Fig. 5a). Specifically, while EPA only accounted for 0.8% of the total FAMEs and DHA was not detectable in Naked Clams grown solely on wood, whereas the respective percentages of EPA and DHA in Naked Clams supplemented with microencapsulated were 1.8% and 1.6% (Fig. 5b). Overall, the total polyunsaturated fatty acid content of Naked Clams supplemented with microencapsulated feed was increased by 75.8% compared to wood only feed (ANOVA, F(1,22) = 6.83, p < 0.05; Tukey–Kramer’s HSD, p < 0.05). The fatty acid profile of Naked Clams grown in this study differed compared to commercially produced blue mussel tissues tested. The dominant fatty acid of the blue mussel tested was arachidonic acid (C20:4n6, 27%) (Fig. 5a), while EPA, DHA accounted for 1.7% and 10.2% of the total FAMEs, respectively. Although EPA levels were similar to Naked Clams supplemented with microcapsules, DHA and total polyunsaturated FAME content (43.9%) exceeded that of Naked Clams.Fig. 5

a Fatty acid methyl ester (FAME) composition (expressed as a fraction of total FAMEs) of Naked Clams fed with wood only (brown), Naked Clams fed with wood + microcapsules (red), and commercially farmed blue mussels (Mytilus edulis) (blue). b EPA (Eicosapentaenoic acid, 20:5(n-3)), DHA (Docosahexaenoic acid, 22:6(n-3)), (c) total saturated, mono-unsaturated (MUFAs) and poly-unsaturated fatty acids (PUFAs) are highlighted. Data are expressed as mean +/− SD of technical replicates (n = 12 for Naked Clams, n = 3 for blue mussels). Letters x, y, and z indicate post-hoc test outcomes, where shared letters indicate no significant difference between groups. Full (empirical and chemical) names of fatty acids are shown in Table S1.

Teredo navalis specimens were sourced from north-eastern US coastal waters. To obtain samples for the experiments, two Eastern pine (Pinus strobus) panels of 200 × 120 × 20 mm were placed in Gloucester Harbour, MA, USA on 4/5/2021, to allow settlement of Teredo navalis larvae. These panels were retrieved on 13/3/22. Larval settlement will have taken place during the warmer summer months of July, August and September in 2021, and so the T. navalis were approximately 9-month-old adults at the point of collection. Blue mussels (Mytilus edulis) for the nutritional analysis were commercially farmed and were sourced from Shetland, Scotland. These mussels were rope-grown, fed on wild marine phytoplankton, and of two years age and market size at the point of harvest in February 2023.

Lipid-walled microcapsules containing 30% powdered Schizochytrium algae by weight were manufactured under patent by BioBullets (BioBullets Ltd, Cambridge, UK). To manufacture the microcapsules a premix slurry containing a waxy encapsulant with antibacterial properties and powdered algae were prepared under conditions of controlled shear. The slurry was pumped into an ultrasonic atomizing nozzle at the top of a cooling chamber. The atomized particles formed near-perfect spheres as they cooled and fell to the chamber base. Further particle cooling was achieved with an air-conveying system before discharge via cyclone to a fluid bed processor. The encapsulated feed was then coated with a proprietary non-ionic surfactant to aid dispersion in water. Further cooling in the fluid bed removed all heat of crystallization from the microcapsules before packaging. All components of the formulation were food grade. The physical characteristics of the microcapsules were quantified using a Malvern Mastersizer particle size analyser. The microcapsules had a mean diameter of 46.6 µm, with a DV(10) of 17.7 µm and DV(90) of 104 µm. The microcapsules had spherical shape, near neutral buoyancy, and were tailored for bivalve consumption and growth^{9,18,19,52,53}.

Laboratory experiments took place in the Davy Building aquarium, University of Plymouth, England, in aerated tanks in temperature-controlled rooms maintained at 15 °C.

For the feed assimilation efficacy experiments, three wooden panels containing T. navalis specimens were placed in three separate 1.6 litre tanks (MW, SDW, and W) from 21/11/2022 till 09/12/2022 (Fig. 1). The panels in tanks MW and SDW had 7 pairs of siphons each, and the panel in tank W 5 pairs of siphons, indicating the presence of 7, 7 and 5 individuals respectively. The panels were exposed to three different dietary regimes – microcapsules + wood (MW), Shellfish Diet 1800 + wood (SDW), and wood only (W). Shellfish Diet 1800 is an algal concentrate blend (Shellfish Diet 1800, Reed Mariculture, California, USA). To ensure valid observations, feeding rates were optimised to minimise pseudofaeces expulsion, given faecal expulsion was the measure of feeding rate in this study. Bivalves are known to expel indigestible particulates in the form of pseudofaeces in high concentrations of particulate matter, and studies on pseudofaeces production have found that particulate concentrations around 2 mg/L show minimal production of pseudofaeces within oysters^55,56. This correlated with using a recommended feeding rate of 3% dw (dry weight) feed per dw bivalve per day⁵⁷. Feeding took place at 09:00 every Monday through Friday, and at 15:00 every Monday through Thursday. Feeding did not occur at 15:00 on Friday as that was when faeces were both counted and collected for weighing. Two half feeds were used in these experiments to avoid the production of pseudofaeces, which could have risen from oversaturation.

For the nutritional profiling experiments, the two wooden panels containing T. navalis juveniles were placed into two separate 10 litre tanks (MW and W) on 09/05/2022 (Fig. 1). The panel in tank MW had 40 pairs of siphons and the panel in tank W 28 pairs of siphons, indicating the presence of 40 and 28 juveniles respectively. The panels were exposed to two different dietary regimes – microcapsules + wood (MW), and wood only (W). Initial juvenile dry weight was measured as 0.069 g, and used to calculate feeding ration for microcapsules at a rate of 3% dry weight (dw) capsule per approx dw bivalve per day⁵⁷. Microcapsules were fed at this rate via a three times weekly feed, with care taken to fully disperse the microcapsules. Juveniles from the MW tank were harvested on 30/06/2022, and juveniles from the W tank were harvested on 02/02/2023.

Faecal rate was observed through two methods, daily digital recording and weekly visual counting. Recording was performed after the 3PM feeding every Monday through Friday using GoPro HERO 11 cameras. For each sample, 10 mins of recording on the siphons was performed for each visible individual in each tank. Recordings were viewed later, and a daily rate was calculated based on the number of faecal pellets seen expelled from the syphons within the timeframe.

The weekly faecal count was observed every Friday at 15:00. High quality top-down pictures of the containers were taken using the GoPros, capturing all the faecal pellets within the container. If one picture of the entire container was not sufficient, smaller pictures were taken instead of each quarter of the tanks. Faecal pellets were counted using visual identification assisted with Microsoft Paint (2023 Version, Microsoft, California, USA), which was used to section off parts of the image and mark counted pellets in an obvious colour, noting the number of pellets counted in each segment. Once these segments were accurately counted and added up, the average rate of each individual was calculated, and the total number would be used for calculating average pellet weight.

Faecal weight was also gathered at 15:00 every Friday. Once pictures of faecal pellets were taken, a syringe was used to collect all the faecal pellets from each sample. These pellets were subsequently filtered from the water using a stand and filter paper, and then washed down with pure water to remove excess salt. The weight of the filter paper was recorded prior to filtering. Following this, the pellets and filter paper were placed in an oven and left over the weekend. After the 09:00 feed on Monday, the dry weight of the pellets was recorded, and by extension the average weight of each pellet from each diet.

Faecal pellets were also collected and pooled from animals under different dietary regimes (wood only, and wood plus microcapsules), and dried in a drying over at 36 °C for 48 h prior to processing for electron microscopy.

For SEM imaging, faecal pellets were applied to a carbon infiltrated tab prior to being sputter coated with approximately 10 nm of gold (Emitech K550, Quorum Technologies, England). Samples were imaged using a JEOL IT800HL Scanning electron microscope (JEOL, Japan) operating in hybrid lens mode. Images were obtained using the secondary electron detector with an accelerating voltage of 2 kV and standard probe current 25. Electron dispersive spectroscopy was carried out using the same instrument utilising an Oxford Instruments Ultim Extreme EDS detector and Oxford Instruments AZtec software (v6.1) (Oxford instruments, England). Samples for EDS were ground using an agate pestle and mortar, before being pressed on to a carbon infiltrated tab and coated with approximately 10 nm of carbon (Q150T, Quorum Technologies, England). EDS maps were collected for 40 min per site at 1024 resolution and 30µs dwell time, using the following working conditions: 6 kV accelerating voltage, analysis probe current 50 (5.7 nA) and 7 mm working distance. Data is normalised by all elements to a 100% total.

For the nutritional profiling, the whole animal tissue from the 40 Naked Clam individuals in the MW tank and 28 in the W tank were homogenised and pooled into 12 samples for each tank type. The following steps of nutritional analyses were then performed on each sample individually.

Frozen bivalve tissues, ~2 g from each sample type (Naked Clams fed with wood only, Naked Clams fed with wood + microcapsules, and commercially farmed blue mussels) were freeze-dried, reduced to a fine powder with bead milling and stored at −80 °C before the further analysis. Shells were removed from the blue mussels, we did not remove the tiny pallets from the Naked Clams. The ratios of different biochemical compounds (proteins, carbohydrates, lipids) were determined using Fourier-transform infrared (FTIR)-attenuated total reflectance (Spectrum Two, PerkinElmer, Germany). Approximately 3–5 mg of finely powdered freeze-dried tissue was pressed on the crystal surface (iATR reflectance cell with a DTGS detector), and scans (wavenumber range of 4000–450 cm⁻¹ at a resolution of 4 cm⁻¹) were recorded and baseline-corrected using Spectrum (version 10, PerkinElmer, Germany). The amide I peak (1624 cm⁻¹), the methyl/methylene peak (2800–3000 cm⁻¹), and the pronounced peak at 1050 cm⁻¹ were used for protein, lipid, and carbohydrate quantification, respectively^58,59. BSA, C-16, and glucose mixed with potassium bromide (KBr) were used as analytical standards. Protein, carbohydrate, and lipid content were then expressed as percentages of dry biomass weight after determining the absolute protein content by a colorimetric method. Powdered biomass (5 mg) was suspended in 5% (v/v) SDS extraction buffer, incubated at 100 °C for 20 min and centrifuged (5000 × g for 5 min). After 1:5 dilution with water, aliquots of the supernatant extracts underwent a 96-well microplate Lowry assay using the Pierce™ Modified Lowry Protein Assay Kit. After the reaction, absorbance at 750 nm was measured in a microplate reader (CLARIOstar Plus, BMG, Germany), and protein content was quantified using a standard BSA curve fitted into a quadratic model as recommended by the manufacturer.

Freeze-dried, powdered bivalve tissues from each of the sample types (~2 mg) were suspended in 1 mL of water and boiled for 20 min. The samples were centrifuged, and the supernatant was used for B₁₂ content measurement. The B₁₂ content was determined by a microbiological bioassay method using a recently evolved B₁₂-dependent strain of Chlamydomonas reinhardtii (metE7⁶⁰). This strain was incubated for 4 days in a TAP-based medium at ~90 µE m⁻² s⁻¹ continuous light, 120 rpm, plus the B₁₂-containing samples, diluted in 24-well plates at a total volume of 2 ml per well. Growth was quantified by measuring optical density at 730 nm. B₁₂ concentration in the samples was estimated by comparing the optical densities obtained to a standard curve of known B₁₂ concentrations (between 8–200 pM B₁₂) fitted with a 4-parameter logistic model (as described in Harrison et al., 2023 under review).

Freeze dried tissues from each sample type were dispersed in an aqueous solution of guanidine and thiourea (6 M/1.5 M); (20% w/v) and diluted with methanol (30%) and TBME (10%)¹⁹. Then lipids were extracted as outlined by Davey et al (2014)⁶¹. Briefly, 200 μL of the dispersions were suspended in 10 mL of (2:1) chloroform and methanol mix, spiked with 200 mL of pentadecanoic acid (C15) (1 mg·mL⁻¹) and sonicated for 30 min on ice. A total lipid extraction was performed as outlined by Davey et al., (2014). Briefly, samples of ~2 mg of freeze-dried biomass were suspended in 10 mL of (2:1) chloroform and methanol mix, spiked with 200 mL of pentadecanoic acid (C15) (1 mg mL⁻¹) and sonicated for 30 min on ice. Then, after addition of 5 mL of water, the tubes were centrifuged at 1000 × g for 3 min at 4 °C to achieve phase separation. The aqueous phase was discarded, the solvent was evaporated from the lower phase with N2 gas (45 °C; GeneVac EZ-2; SP Scientific, Ipswich, United Kingdom), and the extract was then resuspended with 200 mL of n-heptane.

Three 100 mL aliquots of the extract were transferred to a glass tube and trans-esterified with 3 mL of a 2.5% v/v H2SO4/methanol mix, vortexed and placed in a 60 °C water bath for 4 h and then allowed to cool. Following this step, after addition of 3 mL of water and 3 mL of hexane, the tubes were centrifuged at 2000 × g for 3 min to achieve phase separation. The upper hexane phase was then separated, re-extracted and evaporated under the previously defined conditions (Davey et al., 2014). The fatty acid methyl ester (FAME) extracts were resuspended with 120 mL of hexane, separated, and identified using GC (Thermo Scientific Trace GC Ultra) with a Zebron ZB-Wax Capillary GC column (30 m × 0.25 mm, 0.25 µm film thickness, Phenomenex, UK). The injection volume was 1 µL with a 35:1 split ratio with an injector temperature set to 230 °C, using helium as a carrier gas at a constant flow of 1.2 mL·min-1. For the run, the following gradient was used: initial oven temperature 60 °C, 2 min; 150 °C at 15 °C min⁻¹; 230 °C at 3.4 °C min⁻¹. The detector temperature was 250 °C. FAMEs are identified by co-elution with a FAME standard mix (Supelco, 37 Component FAME Mix). The peaks were identified based on the standard mix peak RT and through library search (NIST). The areas were compared as a percentage of the total identified FAMEs.