Biochemical changes throughout early- and middle-stages of embryogenesis in lobsters (Homarus americanus) under different thermal regimes

Lobster source and egg assessment

A total of 15 egg-bearing (ovigerous) lobsters were legally collected (New Hampshire Fish & Game permit, RSA 214:29) in late August and early September (2006) along the New Hampshire (NH) seacoast near Rye, NH and Gunboat Shoals (43°.0274N; 70°.6938W, southern Gulf of Maine) by permitted commercial lobstermen using standard baited traps. Lobsters were transported to the University of New Hampshire (UNH) Coastal Marine Laboratory in Newcastle, NH and lobsters were initially held in a large 1,200 L fiberglass tank with shelters. The holding tank was sourced by ambient sand-filtered seawater (average temp = 15.3 ± 0.5 °C) and was subjected to ambient light. Lobsters in the tank were fed a combination of fresh squid and crabs (Cancer spp.), twice per week.

Lobster carapace lengths (CL) were measured to the nearest 1 mm using digital calipers (Mitutoyo IP 65, Mitutoyo Corp., Japan) (size range: 84–96 mm CL) and a laminated disc tag (diameter = 2.0 cm, Floy Tag Inc., Seattle, WA) was fastened to the propus of each animal with a single ziptie for identification throughout the duration of the study. A subset of eggs from each lobster clutch (n =15–20) were viewed under a dissecting scope on September-5 and staged according to the methods outlined by Helluy & Beltz (1991). Only lobsters whose clutches had eye indices <18% (Perkins, 1972; Goldstein & Watson III, 2015b) were used for this sample to encompass as much of the early development process as possible (Perkins, 1972).

Thermal treatments and sampling

The experimental setup and thermal treatments are described in detail in Goldstein & Watson III (2015b). Briefly, six 0.91 m diameter (600 L) tanks (2 tanks/ treatment) were used to mimic either inshore, offshore, or constant (16 ± 0.4 °C) temperature regimes on a year-round basis. Lobsters in each tank were kept isolated using mesh dividers and each animal received a shelter and was exposed to ambient photoperiod. For the purposes of this study thermal regimes were simulated to either match inshore locations (shallow and coastal, 2–5 km from shore, 8–10 m depth) or the offshore habitats (12–20 km from shore, 20–30 m depth) to which lobsters move to in the fall and overwinter (see Goldstein & Watson III, 2015b). Constant temperatures were chosen to simulate a favorable (and predictable) growth temperature similar to eggs observed in MacKenzie (1988). Temperatures in all tanks were logged every 30 minutes using HOBO pendant loggers (model UA-002-64, Onset Computer, Bourne, MA) and later downloaded using Hoboware software (HOBOware Pro v. 3.0). Inshore temperatures mirrored the ambient seawater that was circulated through the CML seawater system while the temperatures in the offshore tanks were adjusted biweekly to simulate offshore temperatures based on historical and real time data published on the Northeastern Regional Association of Coastal Ocean Observing Systems website (NERACOOS, http://neracoos.org). A total of five ovigerous females were exposed to each of the three temperature treatments and their eggs were sampled at five discrete time periods: twice in the fall (Oct-15 & Nov-15) during initial growth, once in the winter (Jan-15), when eggs remain in a relative stasis, and twice in the spring (Mar-15 & May-25), during periods of rapid growth (Sibert, Ouellet & Brethes, 2004; Goldstein & Watson III, 2015b).

Although in this study each lobster was not truly independently segregated, we chose this design for two reasons: (1) as experimental replicates, lobster eggs are endowed with very thick-layered egg casings that make them isolated from other eggs (with the exception of hatching events when the chorion essentially ruptures, (Talbot & Helluy, 1995); and (2) logistically, we were not able to isolate each lobster on 15 separate incoming seawater lines. This would have been especially problematic for the offshore and constant seawater treatments where we were simulating these conditions with temperature-controlled units. Instead, we created a reservoir tank where we could very accurately control these conditions for each treatment more effectively. Therefore, we attempted to minimize a lack of independence but are confident that keeping females isolated within the same tanks was acceptable given the research question and associated analyses we sought to explore.

At each sampling interval, lobster eggs (∼100/sample) were removed from the center of each lobster clutch with a pair of fine forceps. All egg samples were rinsed and gently agitated with a 0.5% sodium hypochlorite and distilled water solution for ∼1 min., after which they were rinsed with 100% distilled water and blotted dry to remove the cement matrix holding the eggs together (P. Talbot, personal communication, 2011). Previous studies indicated that this chemical separation technique was non-invasive and did not compromise the biochemical integrity of the eggs due to their complex and thickened membranes (Johnson, Goldstein & Watson III, 2011). Each egg sample was divided into 30 egg aliquots then freeze-dried at −40 °C for 24 hr. (Labconco Freeze Dryer 5, Kansas City, MO) and served as a stock for subsequent analyses. Samples were then ground down into a fine powder using an industrial-grade milling machine (Wiley Mill #4, 40 µm mesh screen, Thomas Scientific, Swedesboro, NJ) and samples were stored in labeled polyethelene scintillation storage vials for subsequent analyses.

Biochemical analyses

For each of the five sampling intervals, we determined total lipids and proteins for each lobster in triplicate (n = 30 × 3 egg samples/female) using the methods described in Goldstein (2012). In brief, total protein levels were determined using a modified Lowry method (Lowry et al., 1951) using a BioRad protein assay kit with Coomassie Brilliant Blue G-250 (reagent standard) and bovine serum albumin as a standard (Biorad Laboratories, Hercules, CA). Egg samples were digested in 1N NaOH, filtered, and read on a spectrophotometer (Beckman DU-250; λ = 595). Total lipid was quantified gravimetrically using the general protocol detailed in Bligh & Dyer (1959). The procedure was modified to use a ratio of 1:2:2.5 chloroform-methanol-water extraction, respectively. Samples were dried for 24 hr at 37 °C and stored in a glass desiccator, before being weighed on an analytic balance.

Egg volumes

For calculating egg volumes, 10-15 fresh eggs were removed in the middle of each of four sampling months (Sep, Nov, Jan, and Feb) from each lobster and placed in plastic 2.0 mL storage tubes, preserved in a 4% formalin and sterile seawater solution and stored at 4 °C. Although we had intended to sample and calculate egg volumes in accordance with the same temporal sampling regimes as our biochemical assays, for logistical reasons we were not able to do so. For each egg, a digital picture was taken using a Nikon Coolpix 995 digital camera mounted to a dissecting microsope (Nikon SMZ-2T, Nikon USA Inc., Melville, NY). All egg images were imported into an image processing software (Image J v.1.35, see http://rsb.info.nih.gov/ij/) and a digital measuring tool was used to make calculations of each egg’s longest two axes. All egg calculations were measured to the nearest 0.1 mm (then converted to µm) and values for each sample were averaged (±se). Egg volumes were then calculated using the formula: V = 4∕3∗ (πr³), where r is the radius for spheroid-shaped embryos (see Garcia-Guerrero & Hendrickx, 2004).

Data analysis

Analysis of variance (ANOVA) was used to investigate potential differences in egg protein and lipid content between the three thermal regimes (fixed factor 1) at each of the five sampling intervals (fixed factor 2). A 3 × 5 full factorial design was used and analyzed as a split-plot (SP) ANOVA (whole-plot = temperature, sub-plot = month, df_total = 15) using a PROC MIXED model in SAS v. 9.3 (SAS Institute Inc., Cary, NC). For all ANOVA models, we tested the assumptions of normality, independence, and homogeneity of variance and used a Kuskal Wallis H-test where these assumptions were not supported. Differences between groups were compared using the PDIFF function in SAS. Regression analyses were carried out using JMP v. 9.3 (SAS Institute Inc., Cary, NC) statistical software. All means are expressed ± se.

Water temperatures

Seawater temperatures over the entire course of this study (October-May) averaged 7.1 ± 0.24 °C (range = 2.1–11.2) for inshore laboratory trials, compared with 6.0 ± 0.19 °C (range = 2.8–10.1) for the offshore thermal regime, and 16.2 ± 0.21 °C for the constant treatment tank (Fig. 1). There was an overall significant difference in water temperatures between the constant tank treatment and both inshore and offshore ones (ANOVA, Kruskal-Wallis H-Test; F_2,7 = 10.32, P < 0.0001; Fig. 1) but not between inshore and offshore. It is worth noting that temperatures in the inshore and offshore treatments converged quickly at the outset of the experiment and diverged markedly starting only in early April (Fig. 1). Thus, it is not surprising that there was no difference in embryo development between inshore and offshore treatments, at least until the March sampling, however inshore temperatures were warmer in April and May compared with offshore locations (P < 0.05).

Lipid and protein content

Both total egg lipid and protein levels from inshore and offshore thermal regimes differed from their constant temperature counterpart (SPANOVA; P = 0.0002, Fig. 2). Likewise, the interactive effect of temperature and month was significant for both lipid (F_7,44 = 2.27, P < 0.045) and protein levels (F_7,44 = 46.5, P < 0.0001, Table 1). Overall egg lipid values showed a decrease over time (equation: lipids = 381.76 − 55.00*month, r²_adj = 0.85, P < 0.0001; Fig. 2), falling most dramatically early in the fall (−16.8% inshore, −21.4% offshore, −24.8% constant) and late spring (−63.7% inshore, −59.0% offshore). By contrast, total lobster egg protein values increased over the same time-frame (equation: proteins = −35.53 + 69.11*month, r²_adj = 0.63, P < 0.0001; Fig. 2), and exhibited the largest increases in the fall (60.4% inshore, 57.7% offshore, 66.5% constant) and spring (30.1% inshore, 37.1% offshore).

Egg volumes

Overall, there was a significant increase in egg volume over time for all eggs over all treatments (r²_adj = 0.413, P < 0.001). However, there were no significant changes with respect to egg volume by treatment (F = 0.73, df = 2, P = 0.513) (overall means: inshore = 3,226 ± 163 µm³, offshore = 3,254 ±167 µm³, constant = 3,476 ± 152 µm³), even though there were differences from month-to-month (F = 2.25, df = 3, P < 0.001; Fig. 3).

Lipids and proteins

Many studies conducted on crustacean eggs show that lipids are the major energy reserve (Holland, 1978; Fraser, 1989; Clarke, Brown & Holmes, 1990; Heras, Gonzales-Baro & Pollero, 2000). Although egg yolk lipids were consumed at disparate rates in our thermal treatments and throughout all months, lipids were consumed much more modestly in winter (Fig. 2). This pattern is seen consistently in other crustaceans as well. For example, the egg lipid content of fiddler crab (Uca rapax) decreases significantly (78.4%) through embryogenesis, confirming that lipids constitute an important energy source for embryonic development (Figueiredo et al., 2008). In addition, lipids are also used as structural components of cell membranes that are being formed as they grow (Rosa & Nunes, 2003). Thus, the catabolism of lipids is a classic feature of crustacean eggs and many other crustaceans produce eggs with large lipid reserves that are used throughout embryogenesis (Rosa et al., 2007). Lipid depletion rates are directly related to incubation temperature, and it has been observed in other crustaceans that the energy consumption per day slightly intensified 3 or 4 days before hatching, which could be related to a higher energy production need at this time (Heras, Gonzales-Baro & Pollero, 2000). Yolk lipids tend to become catabolized first followed by yolk proteins. These ratios change and can be used to estimate the cost of egg development at differing temperatures (Sasaki, McDowell Capuzzo & Biesiot, 1986). In the field, lipid profiles (e.g., fatty acids) have been used to identify offshore from inshore lobster eggs (Castell et al., 1995); therefore, it is possible that these constituents are utilized differently across different geographic regions that correspond to contrasting thermal regimes.

For proteins, the consumption rate during embryogenesis may increase as temperature rises (Conceicao et al., 1998). Proteins both function as building blocks for tissues and as a source of energy, when needed (Schmidt-Nielsen, 1991). At elevated temperatures (constant), increases in protein levels were detected and, at these elevated temperatures, tissue synthesis tends to be inefficient and more protein might be used for energy instead (e.g., Garcia-Guerrero, Racotta & Villareal, 2003; García-Echauri & Jeffs, 2018). Therefore, the duration and rates of differing thermal profiles would most certainly affect these biochemical changes and allocations of resource components over time. How this translates to larval survivorship remains poorly understood. However, Sasaki, McDowell Capuzzo & Biesiot (1986) showed that up until Stage IV (i.e., transitionary postlarval stage), lobsters depended, in-part, upon stored lipids and that proper temperature synchronization in these residual lipids maybe favorable to settlement processes.

Egg volume

Increases in egg volume are primarily due to water uptake by the embryo as well as from the retention of metabolic water resulting from respiration (Pandian, 1970; Petersen & Anger, 1997). The associated osmotic changes that ensue during egg development can be an important component to hatching and have also been implicated in mechanically aiding the breakage of the chorion near the time of hatch (Pandian, 1970). Slight changes in lobster egg volume have been previously explained as a function of a plastic response to variations in salinity (Charmantier & Aiken, 1987), and for later eggs, a consequence of physiological factors during development (Pinheiro & Hattori, 2003). In these instances, the movements or residency of lobsters in certain locations where seawater salinities can vary dramatically during certain times of the year (e.g., estuaries; Watson III, Vetrovs & Howell, 1999) may have an impact on aspects of development or hatch, especially near the latter part of egg development (Charmantier & Aiken, 1987).

Female size and condition

In this study we did not specifically address the influence of maternal size or nutritional condition on egg quality in H. americanus. However, other related studies have showed that caloric energy content per egg increases with female size (Attard & Hudon, 1987). Sibert, Ouellet & Brethes (2004) described this relationship by creating a growth index model for egg development and found that larger eggs used yolk lipids more efficiently and sustained faster embryonic growth compared with smaller eggs. An effect of female size on egg reserve allocation has been reported in other decapods including snow crab (Chionoecetes opilio), giant crab (Pseudocarcius gigas), and lobster (Homarus americanus) (Attard & Hudon, 1987; Sainte-Marie, 1993; Gardner, 2001). In lobsters it has been postulated that the effect of female size may mean that larger females make a greater contribution towards egg reserves (Attard & Hudon, 1987). However, the added effect of temperature on egg quality may, in some cases override this effect and maternal nutrition may also modulate egg quality (Goldstein & Shields, 2018). The lecithotrophic nature of lobster eggs is determined largely through the sequestering of maternal nutrients throughout the processes of primary and secondary vitellogenesis during oocyte formation, the latter of which is highly dependent on the female’s organic energy reserves (e.g., lipoprotein; Dehn, Aiken & Waddy, 1983).