Next Article in Journal
Exploring the Failure Mechanism of Container Port Logistics System Based on Multi-Factor Coupling
Previous Article in Journal
A Parametric Study on the Interconnector of Solid Oxide Electrolysis Cells for Co-Electrolysis of Water and Carbon Dioxide
 
 
Search for Articles:
Title / Keyword
Author / Affiliation / Email
Journal
Article Type
 
 
Advanced Search
 
Section
Special Issue
Volume
Issue
Number
Page
 
Journals
JMSE
Volume 11
Issue 5
10.3390/jmse11051064
jmse-logo
Submit to this Journal Review for this Journal Propose a Special Issue
► Article Menu

Article Menu

share Share announcement Help format_quote Cite question_answer Discuss in SciProfiles thumb_up
...
Endorse textsms
...
Comment
first_page
settings
Order Article Reprints
Font Type:
Arial Georgia Verdana
Font Size:
Aa Aa Aa
Line Spacing:
Column Width:
Background:
Article

Diet and Temperature Effects on the Survival of Larval Red Deep-Sea Crabs, Chaceon quinquedens (Smith, 1879), under Laboratory Conditions

by
Nivette M. Pérez-Pérez
1,2,
Matthew Poach
3,
Bradley Stevens
4,
Stacy L. Smith
1 and
Gulnihal Ozbay
1,*
1
Department of Agriculture and Natural Resources, Delaware State University, 1200 North DuPont Highway, Dover, DE 19901, USA
2
Delaware Center for the Inland Bays, 39375 Inlet Road, Rehoboth Beach, DE 19971, USA
3
NOAA NMFS, Milford Laboratory, 212 Rogers Avenue, Milford, CT 06460, USA
4
Department of Natural Resources, University of Maryland Eastern Shore, 11868 College Backbone Road, Princess Anne, MD 21853, USA
*
Author to whom correspondence should be addressed.
J. Mar. Sci. Eng. 2023, 11(5), 1064; https://doi.org/10.3390/jmse11051064
Submission received: 17 January 2023 / Revised: 2 May 2023 / Accepted: 3 May 2023 / Published: 17 May 2023
(This article belongs to the Section Marine Biology)
Browse Figures
Versions Notes

Abstract

:
Declines in commercial crustacean species (such as lobsters, king crab, etc.) have caused an increased interest in the harvest of the red deep-sea crab Chaceon quinquedens. The red deep-sea crab is a federally managed fishery; however, little is known about the species’ general biology, especially the conditions required for larval survival. We aimed to answer two main questions about the life history of the red deep-sea crab. First, is there a common larval hatching pattern between adult female crabs? Specifically, our inquiries are about the duration of the hatching process, daily peak hatching time, and the relationship between female morphometry and the total larvae hatched. Second, which are the factors affecting the survival and development of larval red deep-sea crabs? In order to answer these research questions, we studied the effects of diet (rotifers, Artemia sp., algae, and unfed), temperature (9 °C, 15 °C, and 20 °C), and aquaculture settings. Ovigerous females were obtained from commercial traps and transported to the NOAA James J. Howard Laboratory, NJ. They were placed in the Females Husbandry and Hatching Collection System (FHCS), where the larvae hatched. Hatching of adult females was monitored and measured by volume. A simple linear regression (SLR) was calculated to predict the number of larvae hatched based on the measured volumes, and it was significant (F = 1196; df = 1, 13; R2 = 0.9892, p = 3.498 × 10−14). Duration of hatching period showed an approximate 30 days for adult females red deep-sea crabs, with a common daily maximum hatching time at 22:00 hrs (hatching time seem to follow the sun cycle and the first hours after sunset, Perez, pers. observation). Linear polynomial quadratic regressions were conducted for both years with an interaction term for the two continuous variables (diet and temperature), and were used to model the proportion of larval survival through time. In both years, a highly significant difference was obtained (F = 56.15; df = 4, 2134; R2 = 0.09353; p = < 2.2 × 10−16). There is an effect of diet and temperature in the survival of red deep-sea crabs, but not a combined effect of them.

1. Introduction

The red deep-sea crab, Chaceon quinquedens [1] is a large brachyuran crab from the Geryonidae family that can be found along the continental shelf and slope of the western Atlantic, from Nova Scotia, Canada to the Gulf of Mexico, at depths between 200 and 1800 m [2,3,4]. Red deep-sea crabs (RDSCs) have been economically and commercially important since the 1970s, when a small fishery was established in New England and the Mid-Atlantic states, which has persisted as the oldest fishery of geryonid crabs in the Atlantic [5]. In US waters of the Atlantic Ocean, the RDSC is managed as a single stock with an undetermined minimum legal size, maximum sustainable yield, and overfishing limit due to a lack of critical biological information. However, a fishery management plan (FMP) was established in 2002 by the New England Fishery Management Council (NEFMC) mainly focusing on quota restrictions and fishing effort controls because of the limited biological knowledge available for RDSCs [6] (A total allowable catch was only established in 2002 [7], when the most recent total allowable landing was determined to be 1775 t for the male-only directed fishery [8]. On average, the red crab fishery in the Mid-Atlantic Bight catches about 32 t of live crabs per trip, year-round, in water depths >600 m.
Recently, knowledge of embryonic development and fecundity as well as the physiological and sexual maturity of female RDSCs, which is vital to improving management strategies, has been published [9,10,11]. Physiological and behavioral sexual maturity of red deep-sea crabs were determined using morphological features and ovary and oocyte development to estimate female size at 50% sexual maturity [11]. Physiological size at 50% sexual maturity varied geographically and was estimated to be 61.2 mm carapace length in the Hudson Canyon and 70.8 mm in the Baltimore/Norfolk canyons. Behavioral size at 50% sexual maturity decreased with latitude and was estimated to be 53.9 mm, 62.5 mm, and 65.5 mm carapace length in the Hudson, Baltimore, and Norfolk canyons, respectively [11]. Similarly, Martínez-Rivera and Stevens [10] described the stages and seasonality of red deep-sea crab embryonic development and determined size-specific fecundity. A positive correlation between fecundity and female body size as carapace length was determined via linear regression. The number of eggs per brood ranged from 324,729 to 34,691 for females ranging in size from 106.2 mm to 62.6 mm carapace length [10]. While understanding the reproductive output of female C. quinquedens is critical to fisheries’ management, we must also understand the development of the early life stages of larval red deep-sea crabs. As a commercially harvested species, it is also important to understand the conditions necessary for the survival, growth, and development of target species’ early life stages.
C. quinquedens has a classic Brachyuran cold-water crab life cycle. Brood incubation, zoeal, and larval periods are longer than those for similar species from warm water environments [12]. Larvae usually hatch in spring/summer in shallower waters (200–1800 m) in comparison to their adult counterparts [4]. Larvae are planktonic and develop through six stages: pre-zoea, four zoeal stages, and one megalops stage before juvenile settlement near the mid-slope. This developmental cycle can be completed in 26 days under temperatures ranging from 18 to 21 °C, accelerating development in laboratory conditions where average stage durations were 7 d from zoeal I to zoeal II, 6 d from II to III, 5 d from III to IV, 7 d from IV to megalopa, and finally 14 d from megalopa to first crab [13].
Previous larval development studies under laboratory conditions focused on other red deep-sea crab species, Chaceon granulatus [14] and Geryon trispinosus [15] and were restricted to descriptions of larval development and included no assessment on rearing system, diet, or temperature.
Both temperature and diet affect larval survival and development of planktonic crab species such as C. quinquedens [16,17,18,19,20,21,22]. Larval red deep-sea crabs were reared between 12 and 26 °C [18]. In the coldest temperature treatment (12 °C), developmental abnormalities occurred. In addition, larvae survival to the first zoeal stage was higher at warmer temperatures [18]. The temperature range for optimal larval red deep-sea crab growth was determined to be between 9 °C and 15 °C, as temperatures at 6 °C slowed down development [20]. In addition to temperature effects on larval RDSC development, a lack of suitable prey in sensitive early life cycle phases of Brachyuran crabs can cause death due to starvation even if feeding later resumes [23]. However, comparisons of larval development and survival between RDSCs (Geryon quinquedens) and a marine shallow water crab (Menippe mercenaria) may show some indication that RDSC has some larval nutritional flexibility to survive in a deep-sea environment [19], since there was no difference in the survival of G. quinquedens megalop stage between the rotifer and brine shrimp diets. However, there was evidence of a developmental delay in the larvae reared strictly on the rotifer diet. In contrast, the shallow water crab, M. mercenaria, did not survive to the megalop stage under the rotifer diet [19].
Despite a multi-decade fishery for the red deep-sea crab, little is known about the optimal growing conditions for larval crab stages. Additionally, the effects of the interactions between diet and temperature on larval stages of RDSCs have not been previously assessed. There is interest in the potential viability of the RDSC as a candidate for commercial aquaculture; however, that requires cost-effective high survivability rearing methods similar to those mentioned for red king crabs [24]. These methods also require an understanding of the optimal RDSC larval development conditions, such as the interactions between temperature and diet as well as culture method. Here, we present two options for rearing larval red deep-sea crabs and describe the effects of diet and temperature on larval development for C. quinquedens.
Our objectives are twofold: (i) to determine the optimal conditions for peak survival and development of deep-sea red crab larvae under laboratory conditions that provide a healthy population, and (ii) to study the factors affecting the survival of the early life stages of the red deep-sea crab in aquaculture settings, such as temperature and diet.

2. Materials and Methods

2.1. Crab Collection and Husbandry

Red deep-sea crab (RDSC) ovigerous females were collected by the Atlantic Red Crab Company from sites within Site 3—south of the RDSC distribution in the USA Western Atlantic (Figure 1—MAP; NEFMC 2014).
The fertilized females were transported in a car, using a cooler with ice packs and wet towels, to the NOAA James J. Howard Sandy Hook lab, New Jersey. Alive females were then measured, examined (number of legs, shell condition), and randomly placed in the Female Husbandry and Hatching Collection System (FHCS). The FHCS was designed to maintain live RDSC adult females as well as allow for the ability to collect, count, measure, and transfer hatchlings. The FHCS is a flow-through system (1.2 L/10 min adjustable) with chilled seawater (6.9–11.3 °C, salinity 27–30‰) obtained from the main seawater supply of the NOAA James J. Howard Marine Sciences Laboratory at Sandy Hook, New Jersey. Chilled seawater was UV sterilized and filtered with a combination of a 25 µm and two 5 µm or 50 µm, 20 µm, and 5 µm polypropylene cotton filters (Millipore Sigma, Burlington, MA, USA) depending on seawater sedimentation before entering the individual female buckets. Twelve buckets of 15 L each were distributed and placed within two 210 cm × 60 cm rectangular tanks. Buckets had the capacity to hold one living female red deep-sea crab. Each bucket was equipped with an individual air stone, a water inflow hose, and an outflow extended to the exterior of the bucket. The water exiting through the outflow was directed to a filter mesh placed over a collection cup. This collected the hatched larvae for enumeration. The room containing the system was maintained with a 48 lx (light intensity (luminous flow per unit of area) from 8:30 a.m. (day) to 8:30 p.m. (night) with a 30 min transition to each (sunrise and sunset). One week was used as the acclimation period where handling of animals was minimal. After the acclimation period, adult female crabs were fed every 3 days with defrosted and cut herring fillets to maintain the FHCS and keep hatching containers clean.
During the acclimation period, lost eggs and deposited larvae because of handling were discarded from the collectors. Post-acclimation period, each mesh filter collector was removed and replaced with a clean cup unless no larvae were observed in the filter. All larvae and eggs collected in the collector were transferred with a squirt bottle containing UV-filtered seawater and a funnel to a graduated cylinder (25 mL or 50 mL) until they were settled at the bottom.
Eggs were observed using a dissecting microscope to determine their developmental stage (i.e., eyed, not eyed). Hatched larvae were collected every 24 h, when possible, from all females with advanced eggs. Once eggs were released in larger volumes within a 24 h period, larvae were harvested at 4 h intervals (200, 600, 1000, 1400, 1800, and 2200 h) for 3 consecutive days. The contents of each housing bucket were emptied at each 4 h interval through the larvae collector. If larvae hatching volume decreased substantially after the 3 days, larvae were harvested every 12 h until either (1) hatching volume increased as such that 4 h intervals resumed or (2) hatching volume decreased to less than 1 mL, at which point collection frequency went back to every 24 h. Similar patterns have been shown in other crabs [25] that showed higher hatching at dusk.

2.2. Experimental Rearing System Setups

Two different experiment setups were used, LSFI and LSFII, during the red crab larvae feeding experiments. The experimental setup, LSFI (Figure 2 and Figure 3), consisted of a flow-through system (inflow 12 L/5.1 h) that used filtered seawater (salinity 27‰) from the main seawater supply of the NOAA James J. Howard Marine Sciences Laboratory at Sandy Hook, New Jersey.
During the first experiment, water pumped from Sandy Hook Bay passed through a cyclonic filter followed by sock filters, and finally through sand filtration to a reservoir. From there, the filtered water was then run through a UV sterilizer (Lifegard, Pentair) and two 0.35 μm cartridge filters. To raise the pH to more open ocean conditions, dissolved CO2 was removed by passing the water through two Liqui-Cel® membrane contactors (Membrana-Charlotte, Charlotte, NC, USA) that were run in vacuum mode and with CO2 free air to assist with CO2 stripping. Filtered, sterilized, and CO2-stripped water then proceeded to a head tank, which was bubbled with CO2 free air to ensure against hypoxia before being gravity fed to the inflow line. pH of the inflow water was measured periodically during the first experiment, and it ranged from 7.88 to 8.11 on the total pH scale. Alkalinity of the incoming water was calculated based on pH and dissolved inorganic carbon using CO2 SYS (ORNL/CDIAC-105) [27,28]. Calculated alkalinity ranged from 2137 to 2228 µM/kg of seawater. Salinity of incoming water was about 32 ppt. Inflowing seawater was UV sterilized and treated through a filtration system of chemically resistant pleated polyester filtration 100% cellulose-free cartridges (5 µm, 20 µm, and 50 µm) and then distributed into the experimental buckets. Two tanks containing 10 buckets each (19 L) with lids were used to act as chillers, keeping each set of 10 buckets at 9 °C ± 1 °C and 15 °C ± 1 °C, respectively. These temperatures were selected because they are considered the best range of temperatures for rapid juvenile RDSC growth [19]. Each bucket held 3 smaller containers (750 mL) with a capacity to hold 10 living red deep-sea crab larvae (density 10 larvae/750 mL). Larvae containers were designed to allow the flow of diets through, but not the escape of larvae. Three circular holes (5.5 cm in diameter) were cut, then covered with 150 µm or 250 µm nitex screen mesh (6.8 cm in diameter) depending on the cell size of selected diet. Each temperature treatment had 2 buckets randomly assigned per diet treatment, for a total of 60 larvae per diet type. The light cycle within the tank room was 12 day/12 night hours.
The LSFII system was used in a second experiment which had custom made upwellers composed of a clear conical hatcher (capacity 2.4 L) (Pentair Aquatic Eco-Systems®, Apopka, FL, USA), a top extension with lid, a base reducer of 4-3 inchesPVC (10.16–7.62 cm), a 1 ½ inches PVC (3.81 cm) outflow pipe, and a rectangular aperture covered with a 250 µm mesh (Figure 4). The total capacity of the upweller units with top extensions was 3.15 L, where the fitted conical hatcher could hold 20 living RDSC larvae. Filtered seawater recirculated into the system from the bottom of the upwellers with a flow of 0.25 L/min, through the inflow reservoir 100 and 25 µm mesh, then pumped into the conical hatchers. Two individual circular temperature tanks with chillers were set at 15 °C ± 1 °C and 20 °C ± 1 °C, where each tank contained 12 upwellers. These temperature treatments were selected based on the reported ranges for rapid larvae/juvenile development as well as recent red deep-sea crab surveys, where a temperature of 14.2 °C was recorded at 135 m depth [29]. Each tank had 2 upwellers randomly assigned to a diet treatment, for a total of 60 larvae per diet type. Again, light cycle within the tank room was 12 day/12 night hours. During the second experiment, the water was prepared using distilled water that was enriched and mixed with a commercial sea salt mixture, Instant Ocean Sea Salt for Aquariums, containing every major, minor, and trace element necessary while allowing pH to remain above 8. The sea salt solution we used was meant to mimic natural seawater conditions during the second experiment with consistent water quality conditions. The systems’ water circulations were different: LSFI was flow-through and LSFII was recirculating.

2.3. Larval Crab Feeding Experiments

Both LFSI and LFSII rearing systems were allowed to run for several days prior to the start of the larvae feeding experiments. The mesh collector cups from the FHCS that contained larvae were placed into a cup containing cold UV filtered seawater, so larvae would be suspended and could swim. Actively swimming larvae were collected with a plastic pipette and transferred to small beakers of cold filtered seawater in groups of 10. Each beaker was randomly assigned to both temperature and diet treatments in both the LFSI and LFSII systems. Algal diet culture cultivation and maintenance can be found in supplementary material 1 of the Master’s Thesis of Perez-Perez [26]. Red deep-sea crab larvae were fed diets of rotifers (Brachionus plicatilis), Artemia spp., algae (Tisochrysis lutea, T-Iso), and a mixed combination diet once every morning (24 h cycle) to determine which diet promoted red deep-sea crab larvae growth and development (Table 1). B. plicatilis cultures were started using a live culture obtained from Reed Mariculture Inc. (Campbell, CA, USA). Similarly, Artemia cysts were purchased from Inve Aquaculture (Salt Lake City, UT, USA). Finally, algal cultures of Tisochrysis lutea (T-Iso) were obtained from NEFSC Milford Laboratory (Milford, CT, USA). The LFSI diet experiment included an unfed control treatment (U), a rotifer-only diet (R), a rotifer and Artemia diet (A + R), an Artemia-only diet (A), and finally an Artemia and Tisochrysis lutea (A + AL) diet treatment (Table 1).
In 2016, the LFSII experiment included an unfed control (C), a rotifer-only diet (R), an Artemia-only diet (A), and an Artemia and rotifer diet (A + R) (Table 1). Initial treatments that included Artemia spp. nauplii > 48 h old were too large for red deep-sea crab larvae predation and contributed to their death after 48 h. Therefore, additional diet experiments were used which included unfed (U), a rotifer (R), and T. lutea (AL) groups (Table 1). Diet concentrations varied per diet type and treatment based on larval density (Table 2). Survival rates, as the percentage of larvae alive at a specific time, were recorded daily as well as zoeal stage at death for development over time.

2.4. Statistical Analysis

Survival analysis was calculated as the number of surviving larvae within a given treatment weighted by the number of deaths. To assess the effects of diet, temperature, and zoeal stage on the survival of the red deep-sea crab a linear mixed effects approach was used. Daily recorded larvae mortality was analyzed using linear mixed effects models (LME) from the ‘lme’ package [30] within the R statistical framework (R-4.2.2)). Survival time in days was standardized using the ‘scale’ command within the R statistical framework to convert days into standard deviations from the mean. Linear mixed effects models are an extension of simple linear models to allow both fixed and random effects, and are particularly used when there is non-independence in the data, such as arises from a hierarchical structure, in our case larval survival within each treatment group and across the treatments.
To compare survival differences between LFSI and LFSII systems, only the 15 °C unfed and rotifer treatments from both systems were used. Survival time in days was converted to standard deviations from the mean, and One-Way Analysis of Variance (ANOVA) was conducted using diet (unfed or rotifers) and larval rearing system (LFSI or LFSII) as factors, as well as the interaction between them. All ANOVA statistical analysis was conducted within the R statistical framework (R-4.2.2). A Kruskal–Wallis Test was used to evaluate differences in larval survival within each treatment tank (A–B) and container (I-III) in Larvae Feeding Experimental System 2014 (LFS I). Kruskal–Wallis Test was also used to evaluate differences within each treatment tank’s upwellers (A–C) in Larvae Feeding Experimental System 2016 (LFS II).
In order to obtain the hatched estimation from measure volumes, a simple linear regression with a set intercept on cero was used to describe the relationship between number of larvae and volume and obtained the line formula for estimation. Then, their homogeneity of variances was tested using the Levene’s test, histograms, and a QQ plot. Levene’s test with center = mean was used. Previous Monte Carlo studies indicated that using the mean provided the best power when the underlying data followed a symmetric or moderate-tailed distribution [31].

3. Results

3.1. RDSC Larval Survival

Daily larval survival was calculated as the number of larvae surviving within a given treatment weighted by the number of deaths. In the LFSI system, larvae fed rotifers at both 9 and 15 °C survived the longest, with 20% survival around the 30 day mark (Figure 5). Both the unfed and algae-fed treatments at 9 °C had larvae survive until roughly the 25 day mark, whereas the larvae in the unfed and algae-fed treatments at 15 °C all expired around 20 days (Figure 6). In the LFSII system, larval survival in nearly all diet treatments responded similarly, except for the Artemia spp. at 15 °C fed larval RDSC, which had higher overall survival between days 15 and 18 (Figure 6). Overall, RDSC larvae survived longer in the LFSI system at cooler temperatures, reaching 20% survival at 30 days, whereas 20% survival was hit at the ~17 day mark in the LFSII system. The effects of diet, temperature, and their interactions on survival differed between larval rearing systems and are shown in statistical outcomes (Table 3 and Table 4).
Differences in proportion of survival between diet and temperature groups (Unfed9, Unfed15, Rot9, Rot15, Algae9, and Algae15) were found to be statistically significant using a Kruskal–Wallis Test (Kruskal–Wallis chi-squared: H = 189.13; df = 5, 2139; p = < 2.2 × 10−16). A polynomial quadratic regression with an interaction term for the two continuous variables (diet and temperature) was used to model the proportion of larval survival. A highly statistically significant model was obtained (F = 56.15; df = 4, 2134; R2 = 0.09353; p = < 2.2 × 10−16).
Differences in survival between diet and temperature groups (Unfed15, Unfed20, Rotifers15, Rotifers20, Artemia sp. 15, Artemia sp. 20, Art + Rot15, and Art + Rot20) were found to be statistically significant using a Kruskal–Wallis Test (Kruskal–Wallis chi-squared: H = 22.89; df = 7, 534; p = 0.001781).
A polynomial quadratic regression with an interaction term for the two continuous variables (diet and temperature) was used to model the proportion of larvae that survived. A statistically significant model was obtained (F = 2.531; df = 4, 529; R2 = 0.01136; p = 0.03966).
A Kruskal–Wallis Test was used to evaluate differences within each treatment tank (A–B) and container (I–III) (LFS I), and they were found to be statistically insignificant in 2014, but within the upwellers (A–C) (LFS II), they were found to be statistically significant for most diets in 2016.

3.2. Larval Development

Levene’s test for homogeneity of variances was found to be violated for hatch by group = FemaleID (Levene’s test (center = mean): F = 5.5647; df = 29, 304; p = 1.322 × 10−15) and group = Day (Levene’s test (center = mean): F = 4.5528; df = 52, 281; p = < 2.2 × 10−16). QQ plot and histogram showed a skewed distribution. Owing to these results, an F statistic not assuming homogeneity of variance was calculated.
In the LFSI system, larvae did not develop past the zoeal I stage in the unfed 9 °C and algae 15 °C treatments (Figure 7). Development to the zoeal II stage occurred in the unfed 15 °C, algae 9 °C, and rotifer 9 and 15 °C treatments (Figure 7). Zoeal stage III development only occurred in both rotifer treatments, and development to stage IV only occurred in the 15 °C rotifer treatment (Figure 7). In the LFSII system, all larvae died in the zoeal I stage in both the 15 and 20 °C unfed and rotifer treatments (Figure 8). Larval progression to zoeal stage II and IV was seen in the Artemia spp. 20 °C treatment and only progressed to stage II in the 15 °C treatment (Figure 8). Finally, the combination Artemia spp./rotifer diet in the LFSII system showed larval development to the second and third zoeal stage at both 15 and 20 °C (Figure 8). The effects of diet, temperature, and their interactions on zoeal development differed between larval rearing systems and are shown in Figure 7 and Figure 8.

3.3. Comparison of Survival between the LFSI and LFSII Systems

A comparison between both larval systems was made using only the 15 °C unfed and rotifer treatments in each system (Figure 9). ANOVA results showed significance of the interaction between both diet and rearing system at 15 °C (F = 73.6; df = 3, 296; R2 = 0.4215; p ≤ 0.0001). Larval survival at 15 °C was different between the two rearing systems: Larvae Feeding Experimental System 2014 (LFSI) and Larvae Feeding Experimental System 2016 (LFSII).
For LFS I, daily recorded larvae survival by diet and temperature was tested for homogeneity of variance using a Levene test, histograms, and QQ plots. Homogeneity of variance was found to be violated by the Levene’s test for live larvae proportion by group= Group (Levene’s test (center = median): F = 3.3769; df = 5, 2133; p = 0.004834). Owing to this violated assumption, an F statistic not assuming homogeneity of variance was used. A histogram of larvae survival proportion showed a typical bimodal distribution of the data, and the histogram of larvae survival proportion by days visually presents how larvae died through time. The QQ plot showed a binomial distribution.
For LFS II, daily recorded larvae survival by diet and temperature was tested for homogeneity of variance with a Levene’s test, histogram, and QQ plot. Levene’s test did not show any violation in the homogeneity of variance by group = Group (Levene’s test (center = median): F = 1.2064; df = 7, 526; p = 0.2972), even though the histogram and QQ plot showed a skewed distribution of larval survival proportion.

4. Discussion

This study is the first to analyze the effects of diet and temperature on larval stages of the red deep-sea crab, C. quinquedens, under laboratory conditions. Red deep-sea crab larvae were reared under differing diet and temperature regimes in two separate rearing system experiments. While some individuals did make it to the final zoeal stage, no animals developed to the megalopa stage which may be indicative of delayed development either due to the timing of prey availability or insufficient prey type. Additionally, the length of stage duration may also be indicative of delayed larval development due to diet, temperature, or rearing system. Similar barriers to larval rearing have been extensively reviewed for economically important mud crab species (genus Scylla) [32]. Crab larvae are usually considered carnivorous, however their survival in pelagic ecosystems may be tied to alternative prey resources and the timing at which these prey items are utilized [19,23,32,33]. Feeding behavior and nutritional requirements are different between the zoeal and megalopa stages of several crab species [23]. As such, inappropriate diet choice may decrease the survival of larval crabs as well as delay larval development by increasing stage duration [34,35].
Our results indicate delayed development may have occurred, as nearly all treatments had most larvae still in the zoea I stage even after 20+ days of survival. The developmental time of red deep-sea crabs has been categorized in the past with stage durations ranging from 5 to 14 days between subsequent stages [13]. Therefore, given optimal growing conditions (i.e., prey choice) and/or food availability, C. quinquedens larvae should have progressed through developmental stages within the time frame of our study. Growth to juvenile crab occurred at 18–21 °C with newly hatched Artemia nauplii provided as a prey item [13]. Similarly, Chaceon granulatus was reared successfully to the juvenile stage (after 81–92 d) on Artemia nauplii, where the average stage duration between zoeal I and IV ranged between 8.5 and 12 d, with an average of 46 days spent within the megalopa stage [14]. Survival between stages also varied, where out of 100 individuals 40 made it to the zoeal II stage, 38 to zoeal III, 36 to zoeal IV, 28 individuals to the megalopa stage, and only 2 to juvenile crabs [14].
The importance of diet, including type, size, and timing, in the larval development of several other Brachyuran crab species has also been explored [19,20,21,22,35,36,37,38,39,40,41,42]. Previous studies showed that limited access to prey items (4 to 6 h/day) did not significantly reduce survival of zoea I larvae, but did increase the stage duration for two brachyuran crab species, Cancer pagurus and Chasmagnathus granulata [40]. Additionally, when food limitation continued throughout larval development, survival to metamorphosis was strongly decreased. More recently, the effect of diet during the larval development of shallow-water brachyuran crab species Menippe nodifrons and Callinectes danae was investigated [42]. Larvae from these species were fed with three microalgae species, a rotifer species, and Artemia nauplii, where larvae of C. danae did not reach zoeal III stage on any diet, whereas M. nodifrons larvae were fed enriched rotifers until stage III, after which Artemia nauplii were supplied and they reached the megalopa stage [42]. Both the rotifer and Artemia/rotifer diets in our LFSI and LFSII treatments had larvae that developed to zoeal stage III and IV, whereas the unfed and algae-only diets had high mortality and delayed larval development. Growth, development, and survival of larval Brachyuran crabs Mithraculus sculptus and Mithraculus forceps were evaluated for the aquaculture potential of these ornamental crab species [41]. Starved unfed larvae did not develop past the zoea I stage, while growth rates/development time did not differ between species fed either Artemia or enriched Artemia diets, where mean larval duration was between 8 and 10 d and survivorship to the megalopa stage was over 92% [41].
Developmental delays in larval crustaceans may also be due to differences in essential fatty acid concentrations and the utilization of other biochemical compounds within a particular prey item [34,43,44,45]. Mud crab (Scylla serrata) larvae survival was strongly affected by essential fatty acid (EFA) concentrations between Artemia and enriched rotifer diets, where EFA deficiencies were observed in larvae fed unenriched Artemia [45]. Larvae with these deficiencies had lower survival, longer intermolt periods, and had a narrower carapace when they reached the first crab stage [45]. In the LFSII system, red deep-sea crab larvae were fed with Artemia enriched with DHA Selco (Artemia International), which is rich in highly unsaturated fatty acids. This likely helped larvae in the LFSII system to progress to later zoeal stages. Nghia et al. [44] investigated the effects of docosahexaenoic acid (DHA), eicosapentaenoic acid (EPA), and arachidonic acid (ARA) levels in rotifer and Artemia diets on the survival and development of mud crab larvae, Scylla paramamosain, where survival rate between diet was not statistically significant. However, the DHA/EPA ratio within mud crab larvae prey was a key factor in larval development, where a ~50% HUFA and 0.6 DHA/EPA concentration led to severely delayed/stunted development and metamorphosis failure [43]. In contrast, larvae fed diets with ~30% HUFA and a high DHA/EPA ratio (4) were the best for zoeal development and showed improved metamorphosis rates [43].
In addition to prey type/biochemical composition, discrepancies between larvae and prey size may cause limited larval development and survival [16]. This occurred during our Artemia trial in the LFSI system, where prey items were too big for ingestion by zoea 1 larvae. Increased intake of Artemia as a prey item occurred for mud crab larvae with larval growth, where ingestion of Artemia was low at early zoeal stages (I, II, III), but increased at later stages (IV, V) [46]. However, the small size of rotifers (Brachionus spp. 220–240 µm) likely makes them easy prey for early zoeal stages to capture when compared with larger sized Artemia prey (460–500 µm) [46]. Similarly, Genodepa et al. [47] found that suitable particle size for Scylla larvae during stages I, III, and V were <150 µm, 150–250 µm, 250–600 µm, and 400–600 µm, respectively.
In addition to the variety of diet effects on larval development, temperature has also been documented to impact the reproduction and development of Brachyuran crabs [37,48,49,50,51] Larval stages of brachyuran crabs are the most sensitive to water temperature, and therefore a critical part of crab rearing is assessing how different temperatures affect the growth, survival, and development of larval crabs [51]. In the laboratory, temperature affected the survival and development for Scylla serrata larvae where zoeal stage I was unable to molt to stage II at 17 °C, although normal development until stage V was found to occur at 20–35 °C [48]. In the Dungeness crab, Cancer magister, increased mortality was observed for all larval stages at 20 °C, where 100% mortality occurred at zoeal stage V [38]. Additionally, stage duration varied inversely with temperature, where differences in duration between 10 and 15 °C were greater than those between 15 and 20 °C. For another commercially important crab species, Portunus pelagicus, larval rearing experiments conducted at 30 °C and an ambient range 24–28 °C showed survival to megalopa in a 14 d stage at 30 °C but 100% mortality after day 7 in ambient temperatures. Additionally, survivorship was greater in P. pelagicus larvae at 25 °C and the developmental period was inversely related to temperature [52] It is possible that our temperature regimes were too low for the accelerated growth of larval red deep-sea crabs and, in combination with a poor diet, caused delayed development for most individuals [26].

5. Conclusions

These results provide information on the potential viability of the red deep-sea crab, C. quinquedens, as a commercial aquaculture candidate and indicate delayed larval development due to food choice/limitation/size. Although our red deep-sea crab larvae did not advance to the megalopa stage, we showed that rearing method, temperature, and diet all contribute to the success or failure of larval development in an aquaculture facility. These data provide understanding of potentially improper rearing diets for the red deep-sea crab, C. quinquedens, and whether it is economically viable as an aquaculture species in a modern aquaculture setting. Further research should include refining and improving larval rearing systems, perhaps under food saturation, to better allow for the growth of larvae to the juvenile settlement stage as well as testing other food sources/combinations that are necessary to improve larvae production and development. Larval rearing/development for economically important crab species is still a barrier across taxa [35,41,42]. Therefore, reporting both positive and negative results of larval rearing studies will hopefully benefit those engaged in hatchery research/operations to avoid techniques that are not successful and move toward more viable larval rearing methods.

Author Contributions

Conceptualization, B.S. and G.O.; Methodology, N.M.P.-P.; Formal analysis, N.M.P.-P.; Investigation, N.M.P.-P.; Resources, B.S.; Data curation, N.M.P.-P., B.S. and S.L.S.; Writing—original draft, N.M.P.-P.; Writing—review & editing, M.P. and G.O.; Supervision, G.O.; Project administration, G.O.; Funding acquisition, S.L.S. and G.O. All authors have read and agreed to the published version of the manuscript.

Funding

This research received no external funding.

Data Availability Statement

Data and other information can be obtained from the primary author and the corresponding author.

Acknowledgments

This project was funded by the National Oceanic and Atmospheric Administration (NOAA), the Living Marine Resources Cooperative Science Center (LMRCSC) Grant# NA11SEC4810002, and the NOAA Educational Partnership Program (EPP)/Minority Serving Institutions (MSI) Graduate Research and Training Scholarship Program (GRTSP). Special thanks to Matthew Poach for mentoring the primary author of this manuscript during a graduate internship at the NOAA James J. Howard Marine Sciences Laboratory in Sandy Hook, NJ, as well as all the NOAA James J. Howard Laboratory personnel and students for their support and assistance. Special thanks to Ehren Habeck, Kristen Habeck, DeMond Timmons, and Grant Blank for their technical assistance and guide for every aspect of the experimental setup—the electrical, plumbing, scientific mechanisms, etc. Special thanks to undergraduate students Julianne Murphy and Juliana Tran and high school students Hayley Ehrlich, Charles Lee, and Remi Goddu for their help in the long sampling hours during weekdays and weekends, looking at larvae survival, preparing diets, feeding, and data collection. We would like to thank Ali Parsaeimehr for his assistance with the references section.

Conflicts of Interest

The authors declare no conflict of interest.

References

  1. Smith, S.I. The stalk-eyed crustaceans of the Atlantic coast of North America north of 508 Cape Cod. Trans. Conn. Acad. Arts Sci. 1879, 5, 27–136. [Google Scholar]
  2. Haefner, P.A., Jr.; Musick, J.A. Observations on distribution and abundance of red crabs in Norfolk Canyon and adjacent continental slope. Mar. Fish. Rev. 1974, 36, 31–34. [Google Scholar]
  3. Wigley, R.L.; Theroux, R.B.; Murray, H.E. Deep sea red crab, Geryon quinquedens, Survey off northeastern United States. Mar. Fish. Rev. 1975, 37, 1–21. [Google Scholar]
  4. Steimle, F.W.; Zetlin, C.; Chang, S. Essential Fish Habitat Source Document: Red Deepsea Crab, Chaceon (Geryon) quinquedens, Life History and Habitat Characteristics. In NOAA Technical Memorandum NMFS-NE-163; NOAA: Wood Hole, MA, USA, 2001. [Google Scholar]
  5. Hastie, L.C. Deep-water Geryonid crabs: A continental slope resource. Oceanogr. Mar. Biol. Annu. Rev. 1995, 33, 561–584. [Google Scholar]
  6. Chute, A.; Jacobson, L.; Redding, B. Clarification of Methods Used to Determine Deep Sea Red Crab (Chaceon Quinquedens) Abundance from Data Collected during the 1974 Photographic Survey. In USA Department of Commerce, Northeast Fisheries Science Center Reference Document 13–22; NOAA: Washington, DC, USA, 2013; 19p. [Google Scholar]
  7. Wahle, R.A.; Bergeron, C.E.; Chute, A.S.; Jacobson, L.D.; Chen, Y. The Northwest Atlantic deep-sea red crab (Chaceon quinquedens) population before and after the onset of harvesting. ICES J. Mar. Sci. 2008, 65, 862–872. [Google Scholar] [CrossRef]
  8. NEFMC (New England Fishery Management Council). Atlantic Deep-Sea Red Crab Fishing Years 2014–2016 Specifications, Including A Regulatory Flexibility Analysis; New England Fisheries Management Council, National Oceanic and Atmospheric Administration, National Marine Fisheries Service, Northeast Region: Newburyport, MA, USA, 2016. [Google Scholar]
  9. NEFMC (New England Fishery Management Council). Amendment 3 Fishery Management Plan for Deep-Sea Red Crab (Chaceon Quinquedens) (FMP) Incorporating Specifications for Fishing Years 2011–2013 (March 1, 2011–February 28, 2014) and Including an Environmental Assessment (EA), Regulatory Impact Review (RIR), and Initial Regulatory Flexibility Analysis (IRFA); New England Fishery Management Council, National Marine Fisheries Service: Newburyport, MA, USA, 2014. [Google Scholar]
  10. Martínez-Rivera, S.; Stevens, B.G. Embryonic development and fecundity of the red deep-sea crab Chaceon quinquedens (Smith, 1879) (Decapoda: Brachyura: Geryonidae) in the Mid-Atlantic Bight determined by image analysis. J. Crustac. Biol. 2020, 40, 230–236. [Google Scholar] [CrossRef]
  11. Martínez-Rivera, S.; Long, W.C.; Stevens, B.G. Physiological and behavioral sexual maturity of female red deep-sea crabs Chaceon quinquedens (Smith, 1879) (Decapoda: Brachyura: Geryonidae) in the Mid-Atlantic Bight. J. Crustac. Biol. 2020, 40, 330–340. [Google Scholar] [CrossRef]
  12. Hines, A.H. Larval Patterns in the Life Histories of Brachyuran Crabs (Crustacean, Decapoda, Brachyura). Bull. Mar. Sci. 1986, 39, 444–466. [Google Scholar]
  13. Perkins, H.C. The larval stages of the deep sea red crab, Geryon quinquedens Smith, reared under laboratory conditions (Decapoda: Brachyrhyncha). Fish. Bull. 1973, 71, 69–82. [Google Scholar]
  14. Okamoto, K.; Atsumi, S.; Matsuyama, H. Complete larval development of the deep sea red crab Chaceon granulatus under laboratory conditions. Aquac. Sci. 2005, 53, 93–94. [Google Scholar]
  15. Onishi, K. Larval breeding of a deep-sea crab, Geryon trispinosus (Herbst). Bull. Shizuoka Pref. Fish. Exp. Stn. 1982, 16, 87–95. [Google Scholar]
  16. Sulkin, S.D. The significance of diet in the growth and development of larvae of the blue crab, Callinectes sapidus rathbun, under laboratory conditions. J. Exp. Mar. Biol. Ecol. 1975, 20, 119–135. [Google Scholar] [CrossRef]
  17. Sulkin, S.D. Nutritional requirements during larval development of the portunid crab, Callinectes sapidus Rathbun. J. Exp. Mar. Biol. Ecol. 1978, 34, 29–41. [Google Scholar] [CrossRef]
  18. Rosowski, M.C. The Effect of Temperature on Growth and Dispersal of Larvae of Geryon quinquedens Smith (Brachyura; Geryonidae). Master’s Thesis, University of Delaware, Newark, DE, USA, 1979. [Google Scholar]
  19. Sulkin, S.D.; Van Heukelem, W.F. Ecological and evolutionary significance of nutritional flexibility in planktotrophic larvae of the Deep Sea red crab Geryon quinquedens and the Stone Crab Menippe mercenaria*. Mar. Ecol. Prog. Ser. 1980, 2, 91–95. [Google Scholar] [CrossRef]
  20. Van Heukelem, W.; Christman, M.C.; Epifanio, C.E.; Sulkin, S.D. Growth of Geryon quinquedens (Brachyura: Geryonidae) juveniles in the laboratory. Fish. Bull. 1983, 81, 903–905. [Google Scholar]
  21. Farrelly, C.; Sulkin, S.D. Effect of starvation on megalopal development in the brachyuran crab Hemigrapsus oregonensis. J. Crustacean Biol. 1988, 8, 614–617. [Google Scholar] [CrossRef]
  22. Staton, J.L.; Sulkin, S.D. Nutritional requirements and starvation resistance in larvae of the brachyuran crabs Sesarma cinereum (Bosc) and S. reticulatum (Say). J. Exp. Mar. Biol. Ecol. 1991, 152, 271–284. [Google Scholar] [CrossRef]
  23. Anger, K.; Dawirs, R.R.; Anger, V.; Costlow, J.D. Effects of early starvation periods on zoeal development of brachyuran crabs. Biol. Bull. 1981, 161, 199–212. [Google Scholar] [CrossRef]
  24. Epelbaum, A.B.; Kovatcheva, N.P. Daily food intakes and optimal food concentrations for red king crab (Paralithodes camtschaticus) larvae fed Artemia nauplii under laboratory conditions. Aquac. Nutr. 2005, 11, 455–461. [Google Scholar] [CrossRef]
  25. Stevens, B.G.; Swiney, K.M. Hatch Timing, Incubation Period, and Reproductive Cycle for Captive Primiparous and Multiparous Red King Crab, Paralithodes Camtschaticus. J. Crustac. Biol. 2007, 27, 37–48. [Google Scholar] [CrossRef]
  26. Perez-Perez, N.M. Hatching Periodicity, Survival and Development of the Early Life Stages of Red Deepsea Crab (Chaceon Quinquedens) in Laboratory Conditions: Effects of Diet and Temperature; Natural Resources Graduate Program, Delaware State University: Dover, DE, USA, 2017; p. 116. [Google Scholar]
  27. van Heuven, S.; Pierrot, S.D.; Rae, J.W.B.; Lewis, E.; Wallace, D.W.R. MATLAB Program Developed for CO2 System Calculations. ORNL/CDIAC-105b. Carbon Dioxide Information Analysis Center, Oak Ridge National Laboratory, U.S. Department of Energy: Oak Ridge, TN, USA, 2011. [Google Scholar] [CrossRef]
  28. Lewis, E.; Wallace, D. Program Developed for CO2 System Calculations; Carbon Dioxide Information Analysis Center, Oak Ridge National Laboratory: Oak Ridge, TN, USA, 1998; p. 38. [Google Scholar]
  29. Stevens, B.G.; Guida, V. Depth and temperature distribution, morphometrics, and sex ratios of red deepsea crab (Chaceon quinquedens) at 4 sampling sites in the Mid-Atlantic Bight. Fish. Bull. 2016, 114, 343–359. [Google Scholar] [CrossRef]
  30. Brown, M.B.; Forsythe, A.B. Robust tests for the equality of variances. J. Am. Stat. Assoc. 1974, 69, 364–367. [Google Scholar] [CrossRef]
  31. Hinz, S.; Sulkin, S.; Strom, S.; Testermann, J. Discrimination in ingestion of protistan prey by larval crabs. Mar. Ecol. Prog. Ser. 2001, 222, 155–162. [Google Scholar] [CrossRef]
  32. Waiho, K.; Fazhan, H.; Quinitio, E.T.; Baylon, J.C.; Fujaya, Y.; Azmie, G.; Wu, Q.; Shi, X.; Ikhwanuddin, M.; Ma, H. Larval rearing of mud crab (Scylla): What lies ahead. Aquaculture 2018, 493, 37–50. [Google Scholar] [CrossRef]
  33. Suprayudi, M.A.; Takeuchi, T.; Hamasaki, K. Effects of Artemia enriched with eicosapentaenoic and docosahexaenoic acid on survival and occurrence of molting failure in megalop larvae of the mud crab Scylla serrata. Fish. Sci. 2004, 70, 650–658. [Google Scholar] [CrossRef]
  34. Perez, M.; Sulkin, S. Palatability of autotrophic dinoflagellates to newly hatched larval crabs. Mar. Biol. 2005, 146, 771–780. [Google Scholar] [CrossRef]
  35. Sulkin, S.; Blanco, A.; Chan, J.; Bryant, M. Effects of limiting access to prey on development of first zoeal stage of the brachyuran crabs Cancer magister and Hemigrapsus oregonensis. Mar. Biol. 1998, 131, 515–521. [Google Scholar] [CrossRef]
  36. Sulkin, S.; Lehto, J.; Strom, S.; Hutchinson, D. Nutritional role of protists in the diet of first stage larvae of the Dungeness crab Cancer magister. Mar. Ecol. Prog. Ser. 1998, 169, 237–242. [Google Scholar] [CrossRef]
  37. Sulkin, S.D.; McKeen, G.L. Laboratory study of survival and duration of individual zoeal stages as a function of temperature in the brachyuran crab Cancer magister. Mar. Biol. 1989, 103, 31–37. [Google Scholar] [CrossRef]
  38. Lehto, J.; Sulkin, S.; Strom, S.; Johnson, D. Protists and detrital particles as prey for the first larval stage of the brachyuran crab, Hemigrapsus oregonensis. J. Exp. Mar. Biol. Ecol. 1998, 230, 213–224. [Google Scholar] [CrossRef]
  39. Gimenez, L.; Anger, K. Effects of temporary food limitation on survival and development of brachyuran crab larvae. J. Plankton Res. 2005, 27, 485–494. [Google Scholar] [CrossRef]
  40. Rhyne, A.L.; Penha-Lopes, G.; Lin, J. Growth, development, and survival of larval Mithraculus sculptus (Lamark) and Mithraculus forceps (A. Milne Edwards) (Decapoda: Brachyura: Majidae): Economically important marine ornamental crabs. Aquaculture 2005, 245, 183–191. [Google Scholar] [CrossRef]
  41. Guarizo, M.; Costa, T.M.; Marochi, M.Z. Effect of diet during larval development of Menippe nodifrons Stimpson, 1859 and Callinectes danae Smith, 1869. Aquac. Int. 2020, 28, 1969–1980. [Google Scholar] [CrossRef]
  42. Nghia, T.T.; Wille, M.; Vandendriessche, S.; Vinh, Q.T.; Sorgeloos, P. Influence of highly unsaturated fatty acids in live food on larviculture of mud crab Scylla paramamosain (Estampador 1949). Aquac. Res. 2007, 38, 1512–1528. [Google Scholar] [CrossRef]
  43. Andrés, M.; Estévez, A.; Rotllant, G. Growth, Survival and biochemical composition of spider crab Maja brachydactyla (Balss, 1922) (Decapoda: Majidae) larvae reared under different stocking densities, prey: Larva ratios and diets. Aquaculture 2007, 273, 494–502. [Google Scholar] [CrossRef]
  44. Suprayudi, M.A.; Takeuchi, T.; Hamasaki, K. Cholesterol Effect on Survival and Development of Larval Mud Crab Scylla serrata. HAYATI J. Biosci. 2012, 19, 1–5. [Google Scholar] [CrossRef]
  45. Baylon, J.C.; Bravo, M.E.A.; Maningo, N.C. Ingestion of Brachionus plicatilis and Artemia salina nauplii by mud crab Scylla serrata larvae. Aquac. Res. 2004, 35, 62–70. [Google Scholar] [CrossRef]
  46. Genodepa, J.; Southgate, P.C.; Zeng, C. Diet particle size preference and optimal ration for mud crab, Scylla serrata, larvae fed microbound diets. Aquaculture 2004, 230, 493–505. [Google Scholar] [CrossRef]
  47. Hamasaki, K.; Suprayudi, M.A.; Takeuchi, T. Effect of dietary n-3HUFA on larval morphogenesis and metamorphosis to megalops in the seed production of mud crab, Scylla serrata (Brachyura: Portunidae). Aquac. Sci. 2002, 50, 333–340. [Google Scholar]
  48. Baylon, J.C. Effects of salinity and temperature on survival and development of larvae and juveniles of the mud crab, Scylla serrata (Crustacea: Decapoda: Portunidae). J. World Aquac. Soc. 2010, 41, 858–873. [Google Scholar] [CrossRef]
  49. Gong, J.; Yu, K.; Shu, L.; Ye, H.; Li, S.; Zeng, C. Evaluating the effects of temperature, salinity, starvation and autotomy on molting success, molting interval and expression of ecdysone receptor in early juvenile mud crabs, Scylla paramamosain. J. Exp. Mar. Biol. Ecol. 2015, 464, 11–17. [Google Scholar] [CrossRef]
  50. Azra, M.N.; Aaqillah-Amr, M.A.; Ikhwanuddin, M.; Ma, H.; Waiho, K.; Ostrensky, A.; Tavares, C.P.D.S.; Abol-Munafi, A.B. Effects of climate-induced water temperature changes on the life history of brachyuran crabs. Rev. Aquac. 2020, 12, 1211–1216. [Google Scholar] [CrossRef]
  51. Ikhwanuddin, M.; Azra, M.N.; Talpur, M.A.; Abol-Munafi, A.B.; Shabdin, M.L. Optimal water temperature and salinity for production of blue swimming crab, Portunus pelagicus 1st day juvenile crab. Aquac. Aquar. Conserv. Legis. 2012, 5, 4–8. [Google Scholar]
  52. Bryars, S.R.; Havenhand, J.N. Effects of constant and varying temperatures on the development of blue swimmer crab (Portunus pelagicus) larvae: Laboratory observations and field predictions for temperate coastal waters. J. Exp. Mar. Biol. Ecol. 2006, 329, 218–229. [Google Scholar] [CrossRef]
Jmse 11 01064 g001 550
Figure 1. Designated and used fishing/sampling sites at Newport News by the Atlantic Red Crab Company. Map by Stephanie Martínez-Rivera. The fishing locations are the ones marked with the green dots on the map shown in red oval, provisionally.
Figure 1. Designated and used fishing/sampling sites at Newport News by the Atlantic Red Crab Company. Map by Stephanie Martínez-Rivera. The fishing locations are the ones marked with the green dots on the map shown in red oval, provisionally.
Jmse 11 01064 g001
Jmse 11 01064 g002 550
Figure 2. (A) Diagram of the red deep-sea crab adult female hatching system. (B) Picture of the same hatching system used during the study. Diagram by Dr. Jeffrey Pessutti and picture taken by Tanya Breen in 2014 [26].
Figure 2. (A) Diagram of the red deep-sea crab adult female hatching system. (B) Picture of the same hatching system used during the study. Diagram by Dr. Jeffrey Pessutti and picture taken by Tanya Breen in 2014 [26].
Jmse 11 01064 g002
Jmse 11 01064 g003 550
Figure 3. The LFSI 2014 system had one tank (230 × 60 cm) for each temperature (9 °C and 15 °C), ten buckets in each tank, and three small containers within each bucket in each tank. Each of the three diets were randomly assigned to the buckets and each small container randomly labeled 1 through 3. Each small container held 10 larvae, for a total of 60 larvae per diet per temperature tank.
Figure 3. The LFSI 2014 system had one tank (230 × 60 cm) for each temperature (9 °C and 15 °C), ten buckets in each tank, and three small containers within each bucket in each tank. Each of the three diets were randomly assigned to the buckets and each small container randomly labeled 1 through 3. Each small container held 10 larvae, for a total of 60 larvae per diet per temperature tank.
Jmse 11 01064 g003
Jmse 11 01064 g004 550
Figure 4. The LFSII 2016 system had (A) two circular tanks, one for each temperature (15 °C and 20 °C) and (B) 12 custom made upwellers in each tank. Each of the four diets were randomly assigned to the upwellers and each upweller held 20 larvae, for a total of 60 larvae per diet per temperature tank.
Figure 4. The LFSII 2016 system had (A) two circular tanks, one for each temperature (15 °C and 20 °C) and (B) 12 custom made upwellers in each tank. Each of the four diets were randomly assigned to the upwellers and each upweller held 20 larvae, for a total of 60 larvae per diet per temperature tank.
Jmse 11 01064 g004
Jmse 11 01064 g005 550
Figure 5. RDSC larval percent survival over time for all diet/temperature treatment types in the LFSI system.
Figure 5. RDSC larval percent survival over time for all diet/temperature treatment types in the LFSI system.
Jmse 11 01064 g005
Jmse 11 01064 g006 550
Figure 6. RDSC larval percent survival over time for all diet/temperature treatment types in the LFSII system.
Figure 6. RDSC larval percent survival over time for all diet/temperature treatment types in the LFSII system.
Jmse 11 01064 g006
Jmse 11 01064 g007 550
Figure 7. Larval development stage at death per each treatment in the LFSI system. Data are presented as mean values.
Figure 7. Larval development stage at death per each treatment in the LFSI system. Data are presented as mean values.
Jmse 11 01064 g007
Jmse 11 01064 g008 550
Figure 8. Larval development stage at death per each treatment in the LFSII system. Data are presented as mean values.
Figure 8. Larval development stage at death per each treatment in the LFSII system. Data are presented as mean values.
Jmse 11 01064 g008
Jmse 11 01064 g009 550
Figure 9. Boxplot of square root transformed survival time between diet type rotifer (ROT14 and ROT16) and unfed for each rearing system (LFSI in 2014 and LFSII in 2016) at 15 °C.
Figure 9. Boxplot of square root transformed survival time between diet type rotifer (ROT14 and ROT16) and unfed for each rearing system (LFSI in 2014 and LFSII in 2016) at 15 °C.
Jmse 11 01064 g009
Table
Table 1. Diet treatment combinations for both LFSI and LFSII (2014 and 2016, respectively) with the number of larvae per temperature treatment as well as the number of female broods used to obtain larvae. For 2014, there were two diet setups: the first one is above the line, but the A, A + R, and A + AL diets were not successful, so the diets were changed to the ones shown below the line with double the amount of R and AL. * Unfed (U), Rotifers Brachionus plicatilis (R), Artemia spp. (A), and algae Tisochrysis lutea (AL).
Table 1. Diet treatment combinations for both LFSI and LFSII (2014 and 2016, respectively) with the number of larvae per temperature treatment as well as the number of female broods used to obtain larvae. For 2014, there were two diet setups: the first one is above the line, but the A, A + R, and A + AL diets were not successful, so the diets were changed to the ones shown below the line with double the amount of R and AL. * Unfed (U), Rotifers Brachionus plicatilis (R), Artemia spp. (A), and algae Tisochrysis lutea (AL).
Experiment* DietTemperatureNumber of LarvaeNumber of Broods
2014 LFSI Flow-ThroughU9 °C, 15 °C601
R9 °C, 15 °C601
A9 °C, 15 °C601
A + R9 °C, 15 °C601
A + AL9 °C, 15 °C601
U9 °C, 15 °C601
R9 °C, 15 °C1201
AL9 °C, 15 °C1201
2016 LFSII RecirculatingU15 °C, 20 °C607
R15 °C, 20 °C607
A15 °C, 20 °C607
A + R15 °C, 20 °C607
Table
Table 2. Diet concentrations and volumes fed to the larvae per container ranges for both larval rearing experiments. For 2014, there were two diet setups: the first one is above the line, but the A, A + R, and A + AL diets were not successful, so the diets were changed to the ones shown below the line with double the amount of R and AL.* U—Unfed; R—Rotifers; A—Artemia; A + R—Artemia + Rotifers; A + AL—Artemia + Algae. ** Substituted by an additional rotifer and two algae treatments. Final diet treatments for 2014 are shown below.
Table 2. Diet concentrations and volumes fed to the larvae per container ranges for both larval rearing experiments. For 2014, there were two diet setups: the first one is above the line, but the A, A + R, and A + AL diets were not successful, so the diets were changed to the ones shown below the line with double the amount of R and AL.* U—Unfed; R—Rotifers; A—Artemia; A + R—Artemia + Rotifers; A + AL—Artemia + Algae. ** Substituted by an additional rotifer and two algae treatments. Final diet treatments for 2014 are shown below.
Experiment* DietConcentration
(Number of Diet/mL)		Amount Fed per Container (mL)
2014 LFSI Flow-ThroughU00
R5–340–7
** A520
** A/R2.5/520/range from 40 to 25
** A/AL2.5/10,000,00020/range from 50 to 20
U00
Rrange from 5 to 3range from 40 to 7
AL10,000,000range from 30 to 10
2016 LFSII RecirculatingU00
Rrange from 7 to 4range from 18 to 2
Arange from 0.7 to 0.5range from 7 to 3
A/Rrange from 0.4 to 0.3/from 3.5 to 2.5range from 4 to 1.5/from 9 to 1
Table
Table 3. Kruskal–Wallis Test to evaluate differences in larval survival within each treatment tank (A–B) and container (I–III) in Larvae Feeding Experimental System 2014 (LFS I).
Table 3. Kruskal–Wallis Test to evaluate differences in larval survival within each treatment tank (A–B) and container (I–III) in Larvae Feeding Experimental System 2014 (LFS I).
Treatment Fdfp
Unfed9Containers5.594920.06097
Tanks1.874710.1709
Unfed15Containers3.350620.1873
Tanks0.004786610.9448
Rotifers9Containers3.013820.2216
Tanks6.96371* 0.008318
Rotifers15Containers25.9562* 2.31 × 10−06
Tanks3.87381* 0.04905
Algae9Containers5.542120.0626
Tanks1.097710.2948
Algae15Containers6.54512* 0.03791
Tanks0.06181910.8036
* Statistically significant: p ≤ 0.05.
Table
Table 4. Kruskal–Wallis Test to evaluate differences within each treatment tank’s upwellers (A–C) in Larvae Feeding Experimental System 2016 (LFS II).
Table 4. Kruskal–Wallis Test to evaluate differences within each treatment tank’s upwellers (A–C) in Larvae Feeding Experimental System 2016 (LFS II).
TreatmentFdfp
Unfed153.263420.1956
Unfed2013.9752* 0.00092
Rotifers1510.5632* 0.00509
Rotifers209.03472* 0.01092
Artemia152.597220.2729
Artemia2016.5062* 0.00026
Art + Rot1517.8812* 0.00013
Art + Rot201.513420.4692
* Statistically significant: p ≤ 0.05.
Disclaimer/Publisher’s Note: The statements, opinions and data contained in all publications are solely those of the individual author(s) and contributor(s) and not of MDPI and/or the editor(s). MDPI and/or the editor(s) disclaim responsibility for any injury to people or property resulting from any ideas, methods, instructions or products referred to in the content.

Share and Cite

MDPI and ACS Style

Pérez-Pérez, N.M.; Poach, M.; Stevens, B.; Smith, S.L.; Ozbay, G. Diet and Temperature Effects on the Survival of Larval Red Deep-Sea Crabs, Chaceon quinquedens (Smith, 1879), under Laboratory Conditions. J. Mar. Sci. Eng. 2023, 11, 1064. https://doi.org/10.3390/jmse11051064

AMA Style

Pérez-Pérez NM, Poach M, Stevens B, Smith SL, Ozbay G. Diet and Temperature Effects on the Survival of Larval Red Deep-Sea Crabs, Chaceon quinquedens (Smith, 1879), under Laboratory Conditions. Journal of Marine Science and Engineering. 2023; 11(5):1064. https://doi.org/10.3390/jmse11051064

Chicago/Turabian Style

Pérez-Pérez, Nivette M., Matthew Poach, Bradley Stevens, Stacy L. Smith, and Gulnihal Ozbay. 2023. "Diet and Temperature Effects on the Survival of Larval Red Deep-Sea Crabs, Chaceon quinquedens (Smith, 1879), under Laboratory Conditions" Journal of Marine Science and Engineering 11, no. 5: 1064. https://doi.org/10.3390/jmse11051064

APA Style

Pérez-Pérez, N. M., Poach, M., Stevens, B., Smith, S. L., & Ozbay, G. (2023). Diet and Temperature Effects on the Survival of Larval Red Deep-Sea Crabs, Chaceon quinquedens (Smith, 1879), under Laboratory Conditions. Journal of Marine Science and Engineering, 11(5), 1064. https://doi.org/10.3390/jmse11051064

Note that from the first issue of 2016, this journal uses article numbers instead of page numbers. See further details here.

Article Metrics

Back to TopTop