The research work took place at two distinct facilities: the Mariculture Center of Calheta (CMC; Madeira) and the Experimental Aquaculture Laboratory (AquaLab; Azores). In both facilities, adult specimens of P. aspera and P. candei were captured during the reproductive period (October to April) (Góis et al., 2010; Henriques et al., 2012; Sousa et al., 2017). The specimens collected at CMC facilities were used for assays 1–3, 5–16, and 18, and those collected at AquaLab facilities were used for assays 4 and 17.

A total of 18 assays were realized with the following purposes: assays 1 to 2 deal with “Recollection and management of the broodstock”; assays 3 to 12 deal with “Gametes and larval production methods”; assays 13 to 15 deal with “Larval development and larviculture methods”; assays 16 to 17 deal with “Settlement and metamorphosis in limpets”; and assay 18 deals with “Management of the post-larvae and grow-out”.

The adults dissected for larval production (assays 7–12), larviculture (assays 13–15), settlement (assay 16), and post-larval management (assay 18) were characterized using the average shell length, total mass, and gonadosomatic index ( Supplementary Table 1 for details). The gonadosomatic index was calculated following Sousa et al. (2017): GSI = GM × (TM − SM)^-1 × 100, where GM is the gonadal mass, TM is the total mass, and SM is the shell mass.

For alkaline bath, incubation, and larval culture, we used glass beakers (600 ml) ( Figures 2A, B ) and commercial plastic cups (80–100 ml) ( Figures 2D, E ). For settlement, we used culture cell plates ( Figure 2G ), Petri dishes ( Figure 2H ), and tanks with transparent fiberglass plates ( Figure 2F ). The water employed was filtered (5–20 µm) seawater sterilized with ultraviolet (acronym: FSS).

Larval morphology was used as a criterion to distinguish between viable and deformed larvae, e.g., trochophores (assays 6–15; Figures 9A–C, J–K ) and pediveligers (assays 14–15; Figures 9G–I, L–M ). Larval descriptions are detailed in “Section 3. Larval development and larviculture”.

Assay 1. Impact of physical damage on broodstock survival. The influence of physical injuries on the survival of P. aspera (n = 45) and P. candei (n = 45) in captivity was evaluated. This assay was performed between November 2019 and February 2020. The specimens were tagged (PVC sheets glued with cyanoacrylate; Figures 3D, G, J ). Injuries were categorized into four levels: 0 = no visible injuries, 1 = superficial lesions, 2 = deep lesions, and 3 = evisceration and/or decapitation ( Figures 1H–K ). The specimens were placed in the culture tanks, and mortality was monitored daily to calculate the relative cumulative mortality for each injury level. A bias in this study was that a majority of healthy specimens (level 0; Figure 1N ) were removed prematurely for experimental purposes.

Assay 2. Feeding trials on adult limpets. Acclimated adults of P. aspera and P. candei were placed in plastic baskets located inside the culture tanks. The food was kept for 2 days ( Figures 3G, J ) and tested: commercial fish flakes (ActivPet, Pingo Doce, Portugal) ( Figure 3G ) and artificial meals based on a flour mixture inspired by those developed for Cellana sandwicensis by Nhan (2014) ( Supplementary Table 2 ; Figure 3J ). The feces were examined using a dissecting microscope ( Figures 3H, I, K, L ).

Assay 3. Female fecundity. Female fecundity was calculated as total oocyte production per female in P. aspera (n = 73) and P. candei (n = 37) sampled from October 2020 to April 2021. The shell length was measured, and the gonadosomatic index was calculated. Each female gonad was placed inside a measuring cylinder, leveled up to 100 ml with FSS, and dropped in a beaker. The oocytes from the gonad were extracted ( Figures 5I–J ), and three samples (1 ml volume) were collected and fixed with 300 µl of formaldehyde 37%. The oocytes from the samples were counted to calculate oocyte production.

Assay 4. Spawning induction assays. This assay evaluates different treatments for spawning induction in P. aspera and P. candei. Specimens were collected from intertidal areas in Faial Island (Azores) and acclimatized for 2 weeks in 35-L tanks. Half an hour before the trial, specimens were placed on a tray, left upward, and covered with a wet towel. Trials were performed in an acclimatized room using 500 ml of transparent boxes as experimental containers. Water flow was kept running using filtered (1 µm) seawater (acronym: FSW) at ambient temperature (17.3 ± 0.4°C). All treatments had 8 independent replicates (individual limpets), except for one trial, which was conducted with 5 individuals for each treatment ( Table 1 ). The treatments lasted between 02:00 and 04:30 h or until the first individuals started to spawn. At the end of each trial, the sex ratio was estimated by dissection of 25% of the treated specimens and calculated as: N_male × (N_male + N_female)⁻¹ ( Table 1 ).

The protocol was mainly based on Ferranti et al. (2018) and included five treatments: intense bubbling, UV-light irradiation, hydrogen peroxide (H₂O₂), thermal shock, and a negative control group. The bubbling treatment used specimens placed in a cylindrical colander inside the boxes with FSW and was generated by aquarium air stones positioned under the colander. The UV-light irradiation treatment was tested as it induces spawning in different abalone species (Ebert and Houk, 1984; Moss et al., 1995). The specimens were placed in a flow-through system with UV-treated FSW with a calculated irradiation of 600 mW h L⁻¹. In the thermal shock treatment, the specimens were placed in boxes with FSW whose temperature was increased by 4–5°C. The temperature was maintained at a constant level using a thermostat that was switched off at the end of the treatment. Hydrogen peroxide treatment employed an experimental box filled with FSW in which the pH was increased to 9.1 using 2 M Tris. The specimens were placed in those boxes for 15 min before the addition of 3.2 ml of freshly prepared 10% H₂O₂ solution. The specimens of the control treatment were placed in experimental boxes filled with FSW. At the end of all treatments, seawater was changed in all boxes with FSW and maintained at room temperature until the end of the trial, which lasted 7 ± 1 h.

Assay 5. Measurement of the diameter of oocytes. The oocytes from ten specimens per limpet species (P. aspera and P. candei) were photographed immediately after their extraction using a dissecting microscope (Leica M165, Leica Microsystems, Wetzlar, Germany) connected to a camera and image analysis software (LAS V4.12; Leica Microsystems, Wetzlar, Germany). The oocytes from the same specimens were also measured after receiving an alkaline treatment (NaOH, pH 8.9, 3 h). The minimum and maximum Feret’s diameters were measured for 40 oocytes per specimen using the image analysis software ImageJ 1.53k to calculate the average Feret’s diameter, following Dopchiz et al. (2018).

Assay 6. Variability in female fertility. Larval production (as a ratio of viable trochophores) was analyzed individually in P. aspera (n = 40) and P. candei (n = 16) females sampled from November 2020 to April 2021. Two treatments were applied to the oocytes: control (FSS) and alkaline treatment (NaOH, pH 8.9, 3 h). The specimens were processed in batches of eight females and four males. The males were used to prepare four different sperm pools: ABC, ABD, ACD, and BCD; each letter represents a different male. Each sperm pool was used to fertilize two females. Fertilization was performed at a density of 10⁵ sperm cells ml⁻¹, and incubation lasted 24 h (16 ± 2°C). Three replicates per treatment and per female were realized. Samples were fixed in formaldehyde 6%–8% and identified as viable trochophores (VT; Figures 9A–C ), deformed specimens ( Figures 9J, K ), and unfertilized oocytes ( Figure 5O–S ). The ratio of viable trochophores was calculated as VT × total specimens^-1. Pairwise comparisons of the ratio of viable trochophores between control and alkaline treatments were performed in each female to calculate the percentage of specimens showing significant differences. The influence of the sperm pool was analyzed by pairwise comparisons between females fertilized with the same sperm pool.

Assays 7–8. Alkaline agent (NaOH vs. NH₄OH). This experiment was repeated twice to study the influence of different alkaline treatments on the larval production of P. aspera. The following factors were combined: two alkaline bases (NaOH and NH₄OH), two pH values (8.4 and 8.9), and six times (10, 20, 30, 60, 120 and 180 min), resulting in 24 combined treatments. In addition, a control treatment (FSS, pH 8.0) was applied at the same times specified before (six control treatments). Three replicates per combination were used. The oocytes were rinsed at the end of each corresponding time and kept in FSS. Fertilization was simultaneous for all oocytes (10⁵ sperm cells ml⁻¹). Incubation lasted 24 h (17 ± 1°C). The specimens were fixed and examined to calculate the ratio of viable trochophores, ratio of normal development, and ratio of fertilization (Cañizares et al., 2021).

Assays 9–12. Sperm density. For consistency of the results, evaluation of the optimal sperm density in P. aspera was done in four separate assays, applying six treatments using logarithmic increments of sperm concentration: 10^N sperm cells ml⁻¹; being n = 2, 3, 4, 5, 6, and 7. Four replicates per treatment were used. The oocytes were alkaline treated (NaOH, pH 8.4, 3 h). Incubation lasted 24 h (16 ± 1°C). The specimens were fixed and examined to calculate the ratio of viable trochophores, ratio of normal development, and ratio of fertilization (Cañizares et al., 2021).

Assays 13–15. Validation of the larviculture protocol. Assay 13 compared the viability of the trochophores (24 h post-fertilization) obtained from non-alkalinized oocytes (control using FSS) and from alkalinized oocytes (NaOH, pH 8.9, 3 h). The trochophores were distributed in eight plastic cups per treatment; four cups per treatment were sampled immediately and fixed to calculate (1) the initial density of trochophores and (2) the ratio of viable trochophores. The remaining cups were cultured for 48 h and fixed to calculate (1) the final density of pediveligers and (2) the ratio of viable pediveligers as viable pediveligers × total specimens⁻¹. Student’s t-test (p < 0.05) was used to analyze pairwise comparisons of the ratios of viable larvae and the densities of larvae.

Assays 14 and 15 were performed with oocytes from two different treatments: control (FSS) and alkaline bath (NH₄OH, pH 9, 10 min) and were divided into two sequential parts. First, for each treatment, the oocytes were distributed in four plastic cups (40 ml FSS) and three glass beakers (500 ml FSS) using 100 oocytes ml⁻¹. The beakers were sampled to calculate the initial quantity of oocytes in each beaker. Fertilization used 10⁵ sperm cells ml^−1, and incubation lasted 24 h (16 ± 1°C). The plastic cups were sampled and fixed to calculate the ratio of viable trochophores. In each glass beaker, the upper 400 ml was siphoned, and specimens were gathered using a 55-µm mesh, resuspended in 100 ml of FSS and sampled to calculate the ratio of collected trochophores relative to the initial quantity of oocytes.

At continuation, the collected trochophores from each treatment were pooled and distributed in eight plastic cups (40 ml FSS) and four glass beakers (500 ml FSS) using 4 ± 1 trochophores ml⁻¹. Four cups per treatment were sampled and fixed as described in assay 13. The beakers were sampled to calculate the initial quantity of trochophores per beaker. The remaining cups and the beakers were cultured for 48 h. Then, the plastic cups were sampled and fixed following assay 13. In each glass beaker, the upper 400 ml was siphoned, and specimens were gathered using a 55-µm mesh, resuspended in 50 ml FSS, and sampled to calculate the ratio of collected pediveligers relative to the initial quantity of trochophores.

Assay 16 studied the settlement in P. candei following the methodology described by Castejón et al. (2022). The pediveligers were obtained from alkalinized oocytes (NaOH, pH 8.9, 3 h) and trochophores cultured in glass beakers (12 ± 4 larvae ml⁻¹). The assay was performed by placing pediveligers (72 h post-fertilization) on cell culture plates ( Figure 2G ) using 8 ml of FSS and 4 ± 2 larvae ml⁻¹. The assay lasted 13 days. Six treatments with six replicates each were tested: (1) negative control; (2) shell pieces covered by light pink coralline crusts (143 ± 27 mm²); (3) diatom Halamphora coffeaeformis biofilm; (4) diatom Navicula incerta biofilm; (5) free swimming haptophycean Pavlova sp.; and (6) combined H. coffeaeformis and Pavlova sp. The assay was monitored daily to calculate the ratio of swimming larvae, ratio of crawling larvae, ratio of settled specimens, and ratio of dead specimens. Settlement success was analyzed at Day 13 as the ratio of juveniles (post-larvae with teleoconch).

Assay 17 used gametes of P. candei obtained from a spawning induction in April 2020 (one female and three males; Table 1 ). The oocytes were suitable for fertilization, so alkaline treatment was not required (Nunes et al., 2021). Fertilization was performed using 29 ± 6 oocytes ml⁻¹ and 4.7 × 10⁵ sperm cells ml⁻¹ and lasted 3 h. Pediveligers 72 h post-fertilization were placed in three rectangular tanks (1 L) with FSS at room temperature (17 ± 1°C). The treatments were performed on transparent fiberglass plates (2 × 7 cm; Figure 2F ) in which different biofilms were grown: Amphora sp. (BEA 1588B), Chaetoceros sp. (BEA 0419B), Ulvella lens (CS-801-19), Ulvella leptochaete (BEA 0702B), and Ulva lactuca, in addition to a plate without biofilm as a negative control. The tanks were filled with 400 ml of FSS, placing ca. 200 pediveligers per tank. For 1 week, 50 ml of FSW was added daily to maintain healthy experimental conditions. From Day 8 to Day 64 post-fertilization, the tanks were topped up to 1 L and switched to a flow-through system at constant temperature (17 ± 1°C).

Assay 18. Petri dishes with grown N. incerta biofilms were tested as culture containers for limpet post-larvae. The same algal culture was used as a settlement inducer in both P. aspera and P. candei. The pediveligers (72 h post-fertilization) were obtained from alkalinized oocytes (NH₄OH, pH 9, 10 min). Larvae were cultured in glass beakers (4 ± 1 larvae ml⁻¹). The culture media used for N. incerta was removed before the start of the assay and replaced with 25 ml of FSS. The initial density was 3 ± 1 pediveligers ml⁻¹. The assay was monitored weekly.

All statistical analyses were performed using the statistical software R version 4.1.0. Yuen’s test for trimmed means (package “WRS2 1.1–3”) was used in assays 5–6 as a nonparametric approach for pairwise comparisons (null hypothesis rejected when p < 0.01). The homogeneity of the variances was analyzed using Levene’s test (package “car 3.0-12”), and the normality of the residuals was analyzed using the Shapiro–Wilk normality test (null hypothesis rejected when p < 0.05). The Tukey HSD was used in assays 7–8 to establish the significant differences relative to the treatment with the highest value following Pérez et al. (2016); in assay 6, two combined treatments (NaOH × pH 8.9 × 3 h; NH₄OH × pH 8.4 × 3 h) were removed from the statistical analysis because two replicates from each treatment were accidentally lost. The Tukey HSD was used in assays 9–12 as a parametric method for pairwise comparisons among treatments (null hypothesis rejected when p < 0.05). The Games–Howell test (package “rstatix 0.7.0”) was used in assays 9–10 as an alternative for the Tukey HSD test when the data were normal but not the homogeneity of variances (null hypothesis rejected when p < 0.01). Student’s t-test was used in assays 13–15 as a parametric method to analyze pairwise comparisons (null hypothesis rejected when p < 0.05). Assay 16 analyzed the ratio of juveniles performing the post-hoc Tukey HSD test (null hypothesis rejected when p < 0.05) on arcsine square root transformed data to fit the normality and homoscedasticity assumptions. The HAL+PAV treatment was removed since zero values were obtained in all replicates.

Broodstocks of P. aspera and P. candei were maintained using the same conditions for both species. At the AquaLab facilities (Azores), the adults were kept at a density ranging from 15 to 30 specimens in 55-L cylindrical tanks covered by plastic liners naturally colonized by algae ( Figures 3A, B ). At CMC facilities (Madeira), the adults were kept at a density of 30 to 35 specimens in 200-L cylindrical tanks; in this case, neither plastic liners nor algal growths were employed ( Figures 3C, D ). The environmental conditions were the same at both installations and for both limpet species: flow-through system (1–4 L min⁻¹), at ambient temperature (19 ± 2°C) and salinity (37 ± 1 g L^-1), and under a photoperiod of 12 h light:12 h dark. Aeration was provided to maintain oxygen saturation above 90%. Shallow tanks with flowing water at ambient temperature have also been used to keep different limpets’ species (Hodgson et al., 2007; Ribeiro, 2008; Mau et al., 2018). Other authors preferred recirculation systems (Ferranti et al., 2018; Nakano et al., 2020; Ferranti et al., 2022).

Tank cleaning was performed routinely (every 1–2 days during the first week and every 2–3 days during the second week) to remove feces, mucus, and dead specimens.

The limpets are prone to injuries; consequently, it is suggested to minimize the manipulation as much as possible. If animal manipulations are needed, we suggest to use a blunt knife ( Figure 1G ) to exert a smooth pressure below the shell to detach the specimens, supporting previous studies (Guallart et al., 2013a). The use of plastic liners is another measure that facilitates the detachment of the broodstock ( Figures 3A, B, E ), as suggested by previous studies (Nhan, 2014; Mau et al., 2018; Mau and Jha, 2018). P. aspera and P. candei can move above the upper water level ( Figures 3E, F ). This behavior did not cause associated problems, yet other authors preferred to place artificial turfs at the water level to limit animal movement (Nhan, 2014; Mau et al., 2018; Mau and Jha, 2018) or to generate artificial tides to reduce air exposure (Guallart et al., 2020b; Ferranti et al., 2022).

Broodstock of P. aspera and P. candei were not fed due to the unavailability of diets for these species. However, short feeding trials showed that limpets can accept certain food items (assay 2; Figures 3G, J ). The examination of the feces revealed content related to the commercial fish flakes ( Figures 3H, I ) and the flour-based meals ( Figures 3K, L ). These observations support the limpets as generally opportunistic feeders with an unrestricted diet (Branch, 1971; Della Santina et al., 1993; Burgos-Rubio et al., 2015), which would facilitate the development of diets for aquaculture purposes.

The first limiting factor to start the production of limpets is the availability of mature specimens. Limpets are generally protandrous hermaphrodites in which the smaller and younger specimens tend to mature as males, while the larger and older specimens tend to mature as females (Orton et al., 1956; Wright and Lindberg, 1982; Creese et al., 1990; Espinosa et al., 2006; Lindberg, 2007; Espinosa et al., 2009b; Sousa et al., 2019b). Overfishing affects the sex ratio toward a higher proportion of males and smaller, less productive females (Martins et al., 2017; Sousa et al., 2019b), a factor to be considered when working with specimens from exploited areas. However, the most important factor is the reproductive season. In Azores, the populations of P. candei reproduce during the whole year, peaking in summer (Curdia et al., 2005). In Madeira, the populations P. aspera and P. candei are winter breeders (Góis et al., 2010; Henriques et al., 2012; Sousa et al., 2017), seriously limiting the acquisition of reproductive stocks due to the frequent adverse sea conditions (Torres and Andrade, 2010).

The larval development of P. aspera and P. candei is similar, in terms of both larval morphology and developmental time ( Figure 9 ), except for the body coloration, which resembles that of the oocytes ( Figures 5O–S ). Incubation and larval culture were performed at 17 ± 1°C, and trochophores were observed 17 h post-fertilization. Trochophores showed an oval shape, an apical tuft of cilia, and prototroch cilia ( Figures 9A–C ). The veliger stage was observed 48 h post-fertilization, showing an oval velum and globular protoconch ( Figures 9D–F ). The pediveliger (last larval stage) was observed 72 h post-fertilization, showing eyespots, cephalic tentacles, foot, and operculum ( Figures 9G–I ). The larval morphology of P. aspera and P. candei and the timing of development resemble those described in other limpet species, e.g., Lottia asmi (Kay and Emlet, 2002), L. digitalis (Kay and Emlet, 2002), L. persona (Kolbin and Kulikova, 2011), L. tenuisculpta (Nakano et al., 2020), Patella depressa (Ribeiro, 2008), P. ferruginea (Guallart et al., 2020b; Ferranti et al., 2022), P. ulyssiponensis (Ribeiro, 2008), and P. vulgata (Aquino De Souza et al., 2009).

Settlement and recruitment are key processes in which the planktonic larvae of benthic species must find a suitable site on the benthos to settle and metamorphose to the first post-larval stage (Rodríguez et al., 1993; Jenkins et al., 2009). The loss of the velum, the pediveliger swimming organ, has been used to define the onset of metamorphosis in several marine gastropods (McGee and Targett, 1989; Inestrosa et al., 1992; Searcy-Bernal et al., 1992; Davis, 1994; Roberts and Nicholson, 1997; Gallardo and Sánchez, 2001; Zhao and Qian, 2002; Salas‐Garza et al., 2009; Courtois de Vicose et al., 2010), including different limpet species, e.g., P. aspera (Castejón et al., 2022), P. caerulea (Dodd, 1957; Wanninger et al., 1999), P. ferruginea (Guallart et al., 2020b; Ferranti et al., 2022), and P. vulgata (Dodd, 1957; Wanninger et al., 1999). In limpets, teleoconches are another key characteristic whose presence is restricted to post-larvae (Kay and Emlet, 2002; Ferranti et al., 2018; Nakano et al., 2020; Castejón et al., 2022; Ferranti et al., 2022).

P. aspera and P. candei showed morphological and behavioral similarities during the settlement and metamorphic processes ( Figure 10 ). The pediveligers showed several behaviors: swimming propelled by the velum cilia ( Figures 10 A, E ; Video 4 ), crawling using the foot ( Figures 10B, F ; Video 5 ), withdrawal inside the shell and enclosed by the operculum ( Figures 10C, G ), and resting inside the shell while supported by the foot ( Figures 10D, H ). The early post-larvae is the first post-larval and post-metamorphic stage (Castejón et al., 2022), identified by the loss of the velum and the operculum, and commonly found resting over different surfaces ( Figures 10I, J, L, M ; Video 6 ). The sooner settlement-competency might occur approximately 6 days post-fertilization, since early post-larvae were rarely observed sooner (Castejón et al., 2022). The morphology and behavior of the early post-larvae suggest a distinctive stage involving transformative changes for benthic life, including the development of the digestive structures required to ingest and digest particulate food (Castejón et al., 2022). The next post-larval stage is the juvenile stage (post-larvae with teleoconches), which is reported to be ca. 72 h post-settlement. The juveniles have a wider head, teleoconch, and marked digestive system ( Figures 10K, N ) (Castejón et al., 2022). The juveniles, as active grazers ( Videos 7–8 ), represent the starting point for the grow-out phase.

Traditionally, the settlement process in limpets has been considered a bottleneck associated with high mortality (Corpuz, 1981), with little or no knowledge regarding their requirements and associated settlement cues (Mau and Jha, 2018). Considering the fundamental role of settlement and metamorphosis in the successful development of limpets aquaculture, the search for adequate settlement inducers was a major objective of the present research.

The scientific literature provides scarce information about the management of the early post-larvae and juveniles of limpets, partly due to the limited success reaching these stages (Ferranti et al., 2018; Mau and Jha, 2018; Guallart et al., 2020b; Ferranti et al., 2022). This section resumes the major findings obtained during the present research in relation to the management of the post-larvae using P. aspera and P. candei as model species, which could be helpful for the development of improved protocols.

Post-metamorphic development includes several morphological changes. The head widening is proportionally greater than the animal growth in length ( Figures 10I–N and 13A ). The teleoconch starts as a thin line ( Figure 10J ), which gradually encloses posteriorly as a plane shield surrounding the animal body ( Figures 10K, N ). Body pigmentation was reported approximately 10 days post-settlement ( Figure 13B ), and shell pigmentation was reported 2 months post-settlement ( Figure 13C ). The timing for those developmental changes is more affected by the time after the settlement rather than animal age, i.e., specimens with the same post-fertilization age can show extreme differences if metamorphosis is delayed over time ( Figures 10M, N ).

The juveniles of P. aspera and P. candei showed features and behaviors to be considered for either production or experimental purposes. The widening of the head mentioned before ( Figure 13A ) probably implies the development of the radula and other mouth structures required to actively graze the surroundings, which include shedding of encrusting coralline algae (Castejón et al., 2022) and cultured biofilms ( Figures 13C, D ; Videos 7–8 ), suggesting non-selective grazing, as observed in adults (Branch, 1971; Della Santina et al., 1993; Burgos-Rubio et al., 2015). Several specimens were found to be partially or totally covered by different environmental elements resembling a cryptic strategy to avoid predation ( Figure 13E ; Video 7 ). The juvenile activity was restricted to a 2D environment where the specimens attach tightly and firmly to the surface below. The juveniles showed special affinity for the vertical surfaces, which increases the risk of mortality by desiccation ( Figures 13F, G ).

Direct manipulation of juveniles is not recommended since no reliable methods for animal detachment are available. Moreover, juveniles lying on the back of their shell are unable to recover their normal position ( Figure 13H ; Video 9 ). If individual management is needed, selective pipetting (using a plastic pipette with the tip cut) showed limited success on already detached specimens ( Video 10 ). The juveniles attach firmly to the surface when threatened, so the optimal solution would be to move the surfaces instead of the specimens themselves.

Traditionally, several bottlenecks were present in limpets when considering adult management, gamete release, larviculture and settlement, and metamorphosis induction. The present manuscript represents a significant advance in all these fields, marking a starting point for the aquaculture of different limpet species. Several methodologies were tested, adapted, and optimized for the native species of the Macaronesia region, P. aspera and P. candei, i.e., a methodology for experimental larval production is presented, establishes a larviculture protocol, presents successful settlement inducers, and shows the management and feeding requirements of the post-metamorphic stages. In a few years, it might be possible to close the life cycle of different limpet species and establish large-scale production.

However, researchers have a profuse field to be explored. We acknowledge those authors who previously worked hard on the subject and highlighted many of the specializations and limpets’ requirements. The spawning methods should be polished and optimized; in this sense, the studies realized in P. ferruginea show potential, and the use of hormonal injections should be deeply studied. The determination of female fertility would be important to determine oocyte quality and expected larval productivity. Larviculture methods need to be expanded to allow reliable large-scale production of larvae. The settlement and the grow-out of juveniles are fully experimental, requiring more research before reaching large-scale production levels. In this sense, some of the techniques used for abalone could be adapted for limpets.