The bacterial community composition of American lobster (Homarus americanus) embryos and recently hatched larvae held under different temperature and acidification conditions

The bacterial community composition of American lobster (Homarus americanus) embryos and recently hatched larvae held under different temperature and acidification conditions | Research Square window.SnipcartSettings = { analytics: { enabled: false } }; (function() { var accessVector = localStorage.getItem('access_vector') || ''; window.dataLayer = window.dataLayer || []; if (accessVector) { window.dataLayer.push({ user: { profile: { profileInfo: { snid: accessVector } } } }); } })(); (function(w,d,s,l,i){w[l]=w[l]||[];w[l].push({'gtm.start':new Date().getTime(),event:'gtm.js'});var f=d.getElementsByTagName(s)[0],j=d.createElement(s),dl=l!='dataLayer'?'&l='+l:'';j.async=true;j.src='https://www.googletagmanager.com/gtm.js?id='+i+dl;f.parentNode.insertBefore(j,f);})(window,document,'script','dataLayer','GTM-K279D39R'); Browse Preprints In Review Journals COVID-19 Preprints AJE Video Bytes Research Tools Research Promotion AJE Professional Editing AJE Rubriq About Preprint Platform In Review Editorial Policies Our Team Advisory Board Help Center Sign In Submit a Preprint Cite Share Download PDF Article The bacterial community composition of American lobster (Homarus americanus) embryos and recently hatched larvae held under different temperature and acidification conditions J. Sarah Koshak, Bongkeun Song, Brittany Jellison, Abigail R. Sisti, and 2 more This is a preprint; it has not been peer reviewed by a journal. https://doi.org/ 10.21203/rs.3.rs-6831519/v1 This work is licensed under a CC BY 4.0 License Status: Published Journal Publication published 14 Feb, 2026 Read the published version in Scientific Reports → Version 1 posted 15 You are reading this latest preprint version Abstract Previous research investigating the microbial community of American lobster embryos has long led researchers to believe this habitat comprised only a select few bacterial taxa. However, using 16S rRNA gene sequencing, we show this community to be more diverse than previously thought. We investigated how the bacterial communities of American lobster embryos and larvae change over embryogenesis and hatching in response to two environmental variables. Ovigerous female lobsters caught from Maine and Massachusetts were held under varying temperature and pH regimes that approximated observed and predicted warming and ocean acidification conditions in the Gulf of Maine (GoM) and Southern New England (SNE). The bacterial microbiome associated with the lobster embryos was quantified from two-time points during the experiment, and larvae were collected within 12 hours of hatching. Alpha diversity increased with each life history stage, and embryo and larvae microbiomes shared little community overlap with that in the surrounding tank water. Neither environmental conditions nor lobster origin significantly altered bacterial communities, with life history stage driving alpha and beta diversity. Embryos and larvae shared three core bacterial members identified as members of the genera Rubritalea, Delftia , and Stenotrophomonas . American lobster embryos and larvae appear to have a highly selective microhabitat for bacteria that is not altered by environmental conditions. This leads us to wonder what role the microbiome may have on a developing lobster, and where the microbiome is originating if not from the surrounding seawater. Biological sciences/Microbiology Biological sciences/Microbiology/Communities Biological sciences/Ecology/Microbial ecology microbiome climate change Crustacea metabarcoding Figures Figure 1 Figure 2 Figure 3 Figure 4 Background Crustaceans have received modest attention with respect to their bacterial pathogens and symbionts, and even less attention has been directed at their microbiomes. In general, crustacean embryos and larvae are notoriously susceptible to fungal and water mold (oomycete) infections, most notably those caused by species of Lagenidium, Saprolegnia, and Haliphthoros [ 1 ]. In the early 1990s, a Gram-negative rod-shaped bacterium isolated from lobster embryos was shown to produce a compound with modest anti-fungal properties. This compound was thought to protect the embryos from fungal infections when present in sufficient concentrations [ 2 , 3 ]. Since then, little progress has been made in characterizing the microbial community associated with lobster eggs. Early research focused on culturable bacteria, yielding only a few isolates without taxonomic identifications. As a result, we have a limited understanding of microbial taxa present during one of the lobster’s earliest and most vulnerable life history stages. Given the previous findings of a potential microbial symbiont and the current gap in knowledge, it is important to examine the microbiomes of lobster embryos and larvae using new and advanced techniques. Microorganisms associated with a living host form its microbiome, including bacteria, fungi, viruses, archaea, and protists. Microbiomes of marine animals can originate from the surrounding seawater [ 4 ] or be transmitted across generations, as seen in embryos of some marine sponges [ 5 , 6 , 7 ]. Many of the organisms in egg microbiomes are known to be symbiotic, providing metabolic and immune support to the host, protecting the organism from pathogens, or producing secondary metabolic byproducts that support the host’s fitness [ 8 ]. Conversely, a bloom of pathobionts can slow growth rates, cause pathologies, or compromise host defenses resulting in disease or death to the host [ 9 ]. Given the importance of early life history stages, their importance in recruitment to adult populations, and their sensitivity to climate changes, it is surprising that little research has been conducted on the microbiomes of aquatic and marine embryos and their influence on the developing embryo. Addressing the embryo and larvae microbiome knowledge gap is made more urgent by the impact of climate change. From 1982–2013, sea surface temperatures in the Gulf of Maine warmed by 0.03°C year − 1 , which is three times faster than the global average rate [ 10 , 11 ]. In general, bacteria thrive in a warmer environment with the potential to double their metabolic rate for every 10°C increase in temperature (i.e., the Q 10 ). Some pathogenic species may be able to outpace probiotic species resulting in dysbiosis and disease of their hosts, as observed in American lobsters diagnosed with Epizootic Shell Disease (ESD) [ 12 ]. The initial diagnosis of ESD was hampered by the lack of a comparative baseline of the normal shell microbiome. The disease is now known to be a dysbiosis enhanced by the presence of a few chitinolytic species (e.g., Aquimarina sp.) linked to warming water temperature off Southern New England (SNE) [ 13 , 14 ]. The disease has contributed to the decline of the SNE lobster stock [ 15 , 16 ] and has been identified in lobsters in the Gulf of Maine [ 17 ]. Due to climate change, it is projected to increase in prevalence at a rapid rate in that region [ 18 ]. Although lobster embryos do not appear to be physiologically impacted by ESD [ 19 ], it is prudent to establish baseline data on the embryo microbiome prior to the further influences of warmer water, especially as it may provide insight into potential microorganisms important in abnormal clutches. Embryos are arguably the most sensitive and complex stage in the lobster life cycle and are often found dead or dying in the clutches of egg-bearing females. In fact, pathogenic infection is a leading cause of egg mortality in lobsters and other crustaceans [ 20 , 21 ], but the relationship between mortality and the egg microbiome is not yet known. This study aims to establish a critical baseline for understanding lobster embryo and larvae microbiomes and their potential resilience or vulnerability to environmental changes. Our objectives were, therefore, a) to characterize the microbial community composition of American lobster embryos, b) to evaluate how embryonic stage, temperature, pH, and origin of the lobsters may influence this community, c) to compare the microbiomes of recently hatched lobster larvae to those of embryos and their surrounding tank environment, and d) to identify the core members of microbiomes in different developmental stages of eggs and the first larval stage. Lobster embryos are extruded (i.e., oviposited) into an external clutch on the female’s abdomen and held over a developmental period of 9–11 months. This extended embryogenesis provides a unique opportunity to investigate microbial community dynamics over time. Recent advances in next generation sequencing now allow us to examine the microbial community of embryos and larvae with unprecedented resolution, quantifying the relative abundance of bacterial taxa at different developmental stages. Methods Lobster Collection & Husbandry Ovigerous female lobsters were collected by the Massachusetts Division of Marine Fisheries (courtesy of Dr. Tracy Pugh) and the Maine Department of Marine Fisheries (courtesy of Kathleen Reardon, Special License ME 2019-61-03) as part of their ventless trap surveys. Animals were shipped overnight to the Virginia Institute of Marine Science (VIMS) in October 2020, packed in kelp on top of ice bottles. After a brief period of acclimation in a large, chilled, recirculating tank, where they were held together, 24 animals were separated and randomly distributed among four treatment groups (Supplemental Fig. 1). Lobster pairs (one from ME waters, one from MA waters) were kept in the same aquaria, but physically separated using plastic bins with a mesh top. Briefly, the control treatment group was kept at 10–14°C and pH 8.0–8.1, the warming treatment group was maintained between 14–18°C with pH 8.0–8.1, the acidification treatment group at 10–14°C with a reduced pH 7.5–7.6, and lastly, the multi-stressor treatment group was kept at 14–18°C with a pH 7.5–7.6. Temperatures in all treatment groups were adjusted monthly (hence the range in temperature conditions) to reflect present-day seasonal variation for the southern Gulf of Maine. Each tank contained recirculating seawater (32 psu) that was monitored and controlled autonomously for temperature and pH using a custom system as described in [ 22 ]. Water changes (75%) were performed weekly the day after feeding with a combination of filtered seawater from the York River (Gloucester Point, VA) and artificial seawater adjusted to 32 psu made following the manufacturer’s protocol (Crystal Sea Marine Mix), as described in detail in [ 22 ]. Ovigerous lobsters were fed every 2 weeks with thawed squid during the acclimation period, and then weekly during the treatment period. Sample Collection and Preparation Aliquots of 10–20 eggs were aseptically collected from each lobster using forceps in December 2020 (early group) and March 2021 (late group). Stage I larvae were collected within 12 hours of hatching on various dates. Individual, intact eggs and larvae were flash frozen over liquid nitrogen and stored at -80°C. Eggs and larvae were thawed on ice for quantification of protein content. Briefly, they were homogenized with 200uL of MilliEq water and sonicated at at 20% amplitude for 1-second pulses until no tissue was visible (4–5 pulses). Of the remaining homogenate, 90uL was aseptically removed for determination of protein content, and this procedure is only noted to indicate a freeze-thaw cycle of the homogenized tissue samples. The remaining homogenate was then refrozen and stored at -80°C until DNA extraction for metabarcoding. One egg was used from each ovigerous lobster at each time point. Water samples from each lobster aquarium were collected in 300mL Nalgene bottles near the end point of the experiment in April 2021 and stored at -20° C until thawed for filtration and DNA extraction. To measure bacterial community variation within an egg clutch, two lobsters were selected from which four eggs from each lobster were individually processed as above. DNA Extraction The total DNA was extracted from each remaining tissue homogenate (110 uL, n = 32 embryos) using the DNeasy PowerLyzer PowerSoil Kit (Qiagen) following the manufacturer’s protocol. DNA quality and concentration were measured using Nanodrop (ThermoFisher Scientific) and Qubit (Life Sciences), respectively. The larvae and tank water samples were processed separately but in the same fashion (n = 7 larvae, n = 4 tank water). Water samples were thawed on ice and individually filtered with a Sterivex 0.22 um PES syringe filter (Millipore). Each filter was then removed from the housing, aseptically cut into approximately 2 mm 2 pieces, and the DNA extracted from the filter cuttings as above, following the manufacturer’s protocol. Amplicon library preparation The extracted DNA from embryos was then diluted to a consistent concentration of 2ng/uL for MiSeq PCR, in which indexed, “barcoded” MiSeq 515F-Y forward primers and an 806R reverse primer were used to conduct PCR following the Phusion Hi-Fidelity DNA Polymerase procedure. Initial denaturation was carried out at 98°C for 30s, followed by 30 cycles of 10s denaturation at 98°C, 30s of annealing at 55°C and 30s of extension at 72°C. The final extension was carried out at 72°C for 2 minutes. Amplification was confirmed via gel electrophoresis, and the tagged PCR products (n = 32 embryos) were pooled and homogenized with 10uL of each product. Aliquots of 30uL of the pooled PCR product were electrophoresed using a 2% agarose, and the resulting band excised from the gel and purified using Wizard SV Gel and PCR Clean-up System (Promega). Amplicon length was assessed using the D1000 ScreenTape system (Agilent). The expected fragment size was approximately 300bp. Thirty-four samples, including one positive and negative control, were sequenced using the Miseq Reagent Kits V2 (250bp reads) on the Illumina MiSeq platform at VIMS. The PCR product from a mock community of known composition (ZymoBIOMICS Microbial Community Standard, Zymo Research Corp.) served as a positive control. Extracted DNA from larvae and tank water samples were amplified and sequenced in the same manner as for the embryos on a subsequent occasion (n = 7 larvae, n = 4 tank water). Data Analysis The raw reads were from embryos were trimmed and filtered using the DADA2 analysis package [ 23 ] in the R Studio environment [ 24 ], as separate analyses for each sequencing run, and merged following chimera removal, as per the DADA2 package manager recommendations. Paired-end reads were trimmed by visually assessing quality scores and by using DADA2’s filtering parameters: truncLen = c(200, 160), maxN = 0, truncQ = 2, rm.phix = TRUE, rm.phix = TRUE, and maxEE = 2, and the primers were trimmed using trimLeft = 20. Raw reads from tank water and larvae were filtered using the same parameters, except for truncLen = c(150,150). Sequences were dereplicated and sequence variants inferred using the associated error model. Filtered reads were merged and used to build the amplicon sequence variant tables. Denoised full length sequences were trimmed, and chimeras removed. The sequence table from each run was then merged using mergeSequenceTables() in DADA2. Version 132 of the Silva database [ 25 ] was used to assign taxonomic affiliations. Phyloseq [ 26 ] and ggplot2 packages [ 27 ] in R were used to visualize taxonomic profiles, assess diversity, and perform principal components analysis (PCoA) of the samples. PCoAs were conducted by rarefying samples to the smallest library among the samples and ordinating the distances using Bray-Curtis dissimilarity. All statistical analyses were performed in R, primarily using multivariate community ecology procedures in the base R and vegan [ 28 ] packages. A full record of the DADA2 and phyloseq code used, as well as all statistical analyses are included as Additional files one, two and three. The core community was defined as the genera which occupied 100% of the lobster samples determined by plotting the number of genera shared for each sample type by their prevalence across samples. The core phyloseq objects were agglomerated by genus, using the function tax_glom(ps, taxrank = “Genus”). The microbiome package in R [ 29 ] was used to calculate genus prevalence using the function: core_members(ps, prevalence = .99). The ‘microbiome’ package uses prevalence strictly greater than called, thus, a prevalence of 0.99 was used to determine 100% prevalence of genera among the sample types. The code used to analyze the core is included as Additional file four. Results Embryos The average read depth of the embryos was 338,803 with a minimum of 144,986 and a maximum of 452,715. Developing embryos had diverse bacterial communities consisting of 2,317 ASVs associated with 23 embryo samples, after filtering out organisms with chloroplasts and mitochondria. The negative control contained only one read, and the positive control contained the expected taxa in the mock community. Embryos collected in December (early) tended to have lower Perkin’s Eye Index (PEI) values than those collected in March (late) (Fig. 1 ). Early embryos had lower Shannon diversity (p < 0.05, Kruskal-Wallis test) compared to late embryos (Fig. 1 ). There was no trend observed in early eggs between alpha diversity and PEI, however, alpha diversity increased with PEI in the late samples. The mean Shannon diversity index of early eggs was 2.74 vs. 3.69 for late eggs, with larvae having the highest average index at 4.00. The Simpson and Inverse Simpson index both showed a trend of increasing diversity from early-stage embryos to late-stage embryos to larvae (Supp. table 1). Indices for microbial communities in tank water were slightly above those of early embryos. There were no significant alpha diversity trends noted between the origin of the ovigerous lobster (ME or MA) or among treatment groups (warming or acidification) ( p > 0.05, Kruskal-Wallis test). Also of note, there was no difference in these alpha diversity patterns between rarefied and non-rarefied data, although the observed ASVs were generally lower when rarefied for all samples, as expected (Supp. tables 1 & 2). To better visualize the relative abundance data, the dominant microbial taxa associated with lobster embryos were identified at the Order level. Dominant taxa included Bacteriodales, Chitinophagales, Microtrichales, Rhizobiales, Betaproteobacteriales , Flavobacterales , and Verrucomicrobiales (Fig. 2 ). Initial analyses indicated that one embryo in the early embryogenesis group, T4L1E2, was highly dissimilar to the others, having a disproportionately high relative abundance of Chitinophagaceae , indicating that this embryo may have been unhealthy. It was removed from statistical analyses as an outlier to prevent disproportionate weighting of that sample. Microbiomes on eggs in both early and late stages of embryogenesis grouped differently but they were the closest associates in community composition when analyzed by PCoA (Fig. 3 a) (Bray-Curtis Dissimilarity p < 0.05, PERMANOVA). Microbiomes on early-stage embryos showed a more tightly clustered grouping whereas late-stage embryos had a broader range of diversity demonstrated by a wider spread along axis 1 of the PCoA. No other variables such as treatment group or lobster origin showed significant associations in PCoA or via PERMANOVA ( p > 0.05, Bray-Curtis Dissimilarity). Larvae Larvae from seven lobsters were analyzed from five different tanks with representatives from each treatment group (warming, acidification and multi-stressor) and one control. After initial trimming and filtering, 906 ASVs across the seven samples were identified. The average read depth was 33,008 reads with a minimum of 6,535 and a maximum of 87,975. There were several dominant taxa at the order level consisting of Flavobacteriales, Pseudomonadiales, Bacilliales , and Betaproteobacteriales . Many taxa enriched in the larvae samples and were not associated with embryos or tank water. Microbiomes of larvae had the highest diversity of all sample types with an average Shannon index of 4.00. Notably, the microbiomes of larvae showed distinctly different community compositions from embryos and the tank water samples, even when the larvae were from different tanks (Figs. 3 a & 3 b). In addition, the larvae samples showed no significant trends in community composition in response to the different treatment groups. Tank water Tank water was sampled to examine community differences that might arise from the general milieu of the aquaria in which the lobsters and larvae had been held. Tanks one and two were larvae holding tanks, and tanks three and four were aquaria in which lobsters were housed during the experiment. Tank three was a multi stressor group, and tank four was a control group. The water samples were all collected near the endpoint of the experiment, in April or May. The average read depth across the four samples was 141,405 and after quality filtering, denoising and chimera removal, there remained an average of 103,498 reads per sample. After trimming and filtering, 659 ASVs were detected from tank water samples. At the order level, SAR11 clade, Flavobacteriales , Oceanospirallales and Rhodobacterales dominated the relative abundance of the four samples (Fig. 2 ). The average Shannon index was 2.95. Core community The core community was examined to further understand the stability of taxa present on lobster embryos and larvae and provide a framework for understanding the potential function of the lobster microbiome. We defined the core as the genera present on 100% of the lobster samples (Supplemental Fig. 3). By selecting the genera at 100% prevalence, a distinct core community was identified for each life history stage (Fig. 4 ). Early-stage embryos had 24 core members, with 10 genera found only in this life stage: Bdellovibrio, Blastopirellula, Bradyrhizobium, Caulobacter, Glaciecola, Loktanella, Paucibacter, Oseudoalteromonas, Truepera, and Vibrionimonas . Late-stage embryos had 25 core members, 11 of which were exclusive to this life stage: Aliisedimentitaria, Altererythrobacter, Aquimarina, Aurantivirga, Blastocatella, Candidatus_Nitrotoga, Cocleimonas, Granulosicococcus, Litoreibacter, Maritalea, and Roseovarius . Larvae had 11 core members with seven exclusive to this stage: Acinetobacter, Corynebacterium, Lactobacillus, Lawsonella, Prevotella, Staphylococcus, and Streptococcus . Three genera were shared among all three life history stages; Stenotrophomonas, Rubritalea and Delftia . Early and late stage embryos shared 10 genera: Colwellia, Hellea, Leucothrix, Nitrosomonas, Peredibacter, Pseudrahensia, Pseudomonas, Robiginitomaculum, Sphingorhabdus , Sva0996_marine_group, and Taibaiella. Variation among individual embryos Two lobsters were selected to examine the variation among embryos from the same clutch using four embryos from each lobster, with the resulting microbial compositions analyzed for relative abundance and dissimilarity within and between lobsters. One lobster was from a control group, and the other from a multi-stressor group. All four eggs from each lobster were collected at the same time point. There was 16% variation in beta diversity among the four eggs within an individual, as measured by Bray-Curtis Dissimilarity (Supp. Figure 2 ). In addition, there was no significant difference in diversity between eggs from the same individual when measured by Shannon index (1-sample t-test for outliers, p > 0.05). The limited variation among single embryos from the same clutch confirmed our assumption that the microbial diversity of one embryo is representative of the microbial diversity across an entire clutch was reasonable for this study. Discussion Lobster embryos possess a richly diverse assemblage of bacterial taxa, consisting of 2,561 ASVs. This assemblage changed with embryogenesis, becoming more diverse over developmental time. An increase in Shannon diversity with later developmental stages indicates that the increasing ASV count was not driven by a dominant species type, rather the relative abundances of bacterial taxa were similar among late-stage lobster embryos. Beta-diversity was also driven by life history stage, with early and late embryos harboring distinct communities from larvae and the tank water. All three lobster life history types had uniquely different community structure compared to the surrounding tank water, although communities associated with larvae were more similar to those of the tank water than to other sample types (Figs. 3 a & 3 b). This indicates a tight selection process occurring on lobster eggs, where either the host is exerting control through metabolic influences, or the egg coat is acting as a selective surface for microbial colonization and symbiosis. Surprisingly, no associations were observed based on origin, temperature or pH regimes in this experiment which further implies a selection process early, perhaps during initial oviposition, that is independent of or weakly dependent on the conditions of the external environment. The consistent increase with development in bacterial alpha and beta-diversity observed with lobster embryos may be attributed to several factors. One possibility is the expansion in embryo size during embryogenesis, which nearly doubles the surface area of the embryo [ 30 ], facilitating increased colonization. Another explanation lies in the metabolic waste production and exchange with the external environment throughout embryonic development. The metabolism of lobster embryos slows during the midway point of development, entering a period of diapause during the coldest months where very little, if any, metabolism is occurring [ 31 ]. Then, when water temperatures increase, metabolism increases, which pairs with waste production as increased exchange of small molecules with the external environment [ 32 , 33 ]. The embryos in this experiment did not undergo diapause, as the temperature ranges were higher than typical conditions needed for the dormant phase. There was, however, an increase in oxygen consumption observed throughout embryogenesis [ 22 ], indicating metabolism of the developing embryo rapidly rises in later stages. The increased metabolism of late-stage lobster embryos might act as a chemical signal or additional substrate, attracting distinctive colonizers later in development and leading to a more diverse community. Developmental rates of the lobster embryos from this experiment did not significantly differ among treatment groups [ 22 ]. The implication of this combined work is that not only is the microbiome of lobster embryos resilient to environmental conditions representative of future scenarios of climate change, but the developing larvae within the embryo itself is also seemingly unaffected by these conditions. Finally, members of the early microbiome may act as prey for taxa that colonize older embryos, thereby enriching diversity through shifts in biological relationships within the microbiome itself independent of, or in concert with, the host’s increase in waste products. Change in bacterial community over developmental time is not unique to lobster embryos and was characterized in the hydrothermal vent shrimp, Rimicaris exoculata [ 34 ]. However, this system is quite different as mineral deposits on the egg coat of the shrimp also increased with development due to their proximity to hydrothermal vents, providing an alternative colonization substrate. Moreover, these communities are heavily influenced by extreme vent fluid biogeochemistry and maternal aeration in later development. Lobsters in this study were held in artificial seawater with varying water temperature and chemistry indicating the microbiomes on lobster eggs are resilient to environmental conditions within this system, and a change in metabolism from the lobster embryo is likely driving the observed changes in microbiome structure. Microbiomes of recently hatched lobster larvae had higher alpha and distinct beta diversity when compared to those of embryos and tank water, perhaps due to the development of the larval gut microbiome upon hatching. No studies have described the gut microbiome in larval, juvenile, or adult American lobsters. One study examining the larval gut microbiome of the European lobster ( Homarus Gammarus ) reported a much less diverse assemblage than was found on embryos and larvae in this study, reporting an average Shannon index of around two (exact mean not reported, n = 183) [ 35 ]. Nonetheless, several of the bacteria identified in the larval samples of this study are known gut colonizers in other species. For example, Bacilli are known gastrointestinal probionts in many organisms, and have been shown to increase gut diversity in the farmed shrimp Penaeus vannamei when administered with Streptomyces [ 36 ]. The European lobster, H. Gammarus , larvae showed improved growth rates when fed a Bacilli probiotic [ 37 ]. Beneficial Clostridia have been found in the gut of farm raised the freshwater prawn, Macrobrachium rosenbergii [ 38 ]. The microbial community of the tank milieu had lower alpha-diversity compared to those of the lobsters at every stage of development examined and exhibited the most overlap with the larval community. Nonetheless, when microbiomes of larvae and tank water were analyzed independently, beta-diversity was distinct between the sample types indicating the cuticle of the larva is selective for bacteria absent in the seawater, and/or that the gut microbiota is enriched by the nutrients the larvae are filtering from the seawater (the larvae were not fed during this time). These results support previous findings where the diversity of microbes in larvae of the swimming crab ( Portunus trituberculatus) were shown to be unique from the rearing tank water [ 39 ] and further supports the selective nature of the decapod shell, be it egg or cuticle. We define core members of a microbiome as those simply more prevalent on their hosts than other constituents. They may or may not be beneficial. Embryos and larvae had a diverse core community that was consistent with complex carbon degrading (saprobic) organisms such as those from the families Chitinophagaceae and Flavobacteriaceae , and genus Leucothrix . Leucothrix is particularly interesting because it is a known marine fouling organism, often invading crustacean and fish embryos [ 40 ], and marine alga [ 41 ]. Leucothrix was identified as a core member in both early and late stage embryos but was found in higher abundance in the early stage. Perhaps the community of early-stage eggs is more susceptible to Leucothrix infiltration given the lower overall microbial diversity at this stage. Another core community member in late embryo samples belonged to the family Flavobacteriaceae , which comprised 263 ASVs among the samples. Members of this family are common colonizers of marine invertebrates, fish, and alga. Previous genomic analysis in the marine clade of this family revealed several genes encoding polymer-degrading enzymes for complex carbon metabolism, and a high prevalence of genes encoding secondary metabolites such as antioxidants and cytotoxic compounds [ 42 ]. The genes found in the marine clade of Flavobacteriaceae may be important for late-stage lobsters in preventing the overgrowth of fouling organisms susceptible to cytotoxic defenses (i.e. Leucothrix ). Bacteria of the genus Aquimarina are members of the Flavobacteriaceae family and were associated with only late-stage embryos. The overgrowth of Aquimarina spp . leads to necrotic shell lesions associated with the progression of Epizootic Shell Disease of the juvenile and adult American lobster, particularly in warming waters [ 13 ]. The presence of this bacteria on the embryo of lobster, and from adult lobsters without any indication of disease, supports previous research postulating that Aquimarina is typically a non-pathogenic colonizer that can outcompete other species of the carapace microbiome when external conditions are altered (i.e. when water temperature is elevated) [ 14 , 13 , 43 ]. Delftia, Stenotrophomonas, and Rubritalea were core members to all life history stages. Delftia have recently been identified as potential bioremediators, with some marine species able to degrade napthalene [ 44 , 45 ], heavy metals [ 46 ], and polycyclic aromatic hydrocarbons (i.e. forever chemicals known as PAHs) [ 47 ]. The role Delftia play on lobster embryos are larvae is likely related to carbon degradation from other sources. Delftia and Stenotrophomonas have been identified as a core larval member in the hard-shelled mussel ( Mytilus coruscus ) and was shown to decrease in abundance when experimentally exposed to increased seawater temperature [ 48 ]. Stenotrophomonas spp. are known to be important in nutrient cycling, particularly with nitrogen and sulfur [ 49 ]. Rubritalea was the final core genus shared between larvae and all embryos, which has previously been identified in ascidia (sea squirt) [ 50 ], deep-sea seawater [ 51 ], various sponges [ 52 ], and a marine gastropod [ 53 ]. Not much is known about this genus; however, some species are able to produce secondary metabolites, reduce nitrate, and are red in color due to the carotenoids they contain [ 52 , 54 ]. On sponges and seaweeds, they are thought to play a symbiotic role by scavenging reactive oxygen species or producing antioxidants (55, 56, 57). Rubritalea may play a similar role on developing lobster embryos. Conclusion We show that lobster eggs maintain a unique and diverse assemblage of bacteria over the course of development. The diversity of this community increases over time, and the eggs share little overlap with the surrounding tank water. Larvae have distinct communities from late-stage embryos, and harbor several unique taxa not detected in the environment around them. In the context of the initial experiment examining warming and acidification, neither the core communities nor the broader microbial communities changed in response to these variables. That is, the bacterial microbiome appeared resilient to changes in temperature and acidification. Our study shows that lobster embryos offer selective and stable microhabitats with which to address questions about the origin of these microbiomes, and the need to examine their longevity and change over the span of embryogenesis in finer detail. Abbreviations GoM Gulf of Maine SNE Southern New England ESD Epizootic Shell Disease PCoA Principal Component Analysis CI Confidence Interval ME Maine MA Massachusetts DMR Department of Marine Resources ASV Amplicon Sequence Variant VIMS Virginia Institute of Marine Science DNA Deoxyribonucleic Acid PAH Polycyclic aromatic hydrocarbon PCR Polymerase Chain Reaction rRNA Ribosomal Ribonucleic Acid Declarations Availability of data and materials The datasets supporting the conclusions of this article are available at the National Center for Biotechnology Information (NCBI) BioProject repository, accession number PRJNA1227997 (https://www.ncbi.nlm.nih.gov/sra/PRJNA1227997). Data and R code are accessible at https://figshare.com/s/d052cebd1a2ef3f6cb5b. Competing Interests The authors declare that they have no competing interests. Funding We thank the following for partial funding of this project: NOAA National Sea Grant’s American Lobster Initiative #NA19OAR4170393 and #NA24OARX417C0578 as well as the Batten School for Coastal and Marine Sciences, Virginia Institute of Marine Science, College of William and Mary. Authors Contributions JSK conceived the study, performed 16S rRNA gene laboratory assays, analyzed the 16S rRNA gene sequencing data, wrote the manuscript, and prepared figures. BS contributed resources and training, and helped with 16S rRNA data analysis. BJ, ARS and ER conceived the climate change study, contributed to animal husbandry, and collected samples. JDS conceived the climate change study, collected samples, wrote the manuscript, and facilitated the project. All authors edited the manuscript and approved the final draft. Acknowledgements : We would like to thank Drs. J. Goldstein, A. R. Wargo, and K. Reece for their helpful critiques of the manuscript. We are indebted to Kathleen Reardon and her team at the DMR in Maine, and Dr. Tracy Pugh and her team at DMF, Massachusetts for acquiring and shipping lobsters to us. Brett Sweezey and Gabe Thompson helped with lobster maintenance over the course of the experiment. Mara Walters and Lilly Blume contributed by helping with DNA extractions, PCR, helpful discussions, and various other assistance in the lab. This work was funded by the NOAA American Lobster Initiative #NA190AR4170393 to ABR and JDS. References Shields, Jeffrey D., Fran J. Stephens, and Brian Jones. "Pathogens, parasites and other symbionts." Lobsters: biology, management, aquaculture and fisheries” (2006): 146-204. Gil-Turnes, M. S., Hay, M. E., & Fenical, W. (1989). Symbiotic marine bacteria chemically defend crustacean embryos from a pathogenic fungus. Science , 246 (4926), 116-118. Gil-Turnes, M. S., & Fenical, W. (1992). Embryos of Homarus americanus are protected by epibiotic bacteria. 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13:26:18","extension":"html","order_by":15,"title":"","display":"","copyAsset":false,"role":"acdc-reference","size":138732,"visible":true,"origin":"","legend":"","description":"","filename":"earlyproof.html","url":"https://assets-eu.researchsquare.com/files/rs-6831519/v1/464641af43a8eb77b7a48bfd.html"},{"id":92953148,"identity":"aa01b88b-2f81-425e-b9c9-6a0cfdb21bed","added_by":"auto","created_at":"2025-10-07 13:34:18","extension":"png","order_by":1,"title":"Figure 1","display":"","copyAsset":false,"role":"figure","size":165268,"visible":true,"origin":"","legend":"\u003cp\u003eShannon Diversity for each embryo sample shown as a function of the Perkin’s Eye Index. There is no relationship between PEI and alpha diversity in early embryos. In late embryos, there is a positive correlation between PEI and Shannon diversity indicating that as embryos enter later stages of development, they tend to recruit more diverse bacteria to the surface of the egg.\u003c/p\u003e","description":"","filename":"1.png","url":"https://assets-eu.researchsquare.com/files/rs-6831519/v1/b222c5f88338869482731cf9.png"},{"id":92952908,"identity":"4dac0328-e85d-41f3-b2d3-7d6db9fd3784","added_by":"auto","created_at":"2025-10-07 13:26:17","extension":"png","order_by":2,"title":"Figure 2","display":"","copyAsset":false,"role":"figure","size":562084,"visible":true,"origin":"","legend":"\u003cp\u003eRelative abundance of bacteria at the order level by sample type. A general trend of increasing richness is observed in early vs. late-stage lobster embryos. The rearing tanks have notably fewer taxa at the order level, and several taxa associated with lobsters were not found in the tank milieu.\u003c/p\u003e","description":"","filename":"2.png","url":"https://assets-eu.researchsquare.com/files/rs-6831519/v1/e70ede99eeea61d1f10e5a31.png"},{"id":92954256,"identity":"76a63158-c342-44bb-8e4f-3c39e34d5a64","added_by":"auto","created_at":"2025-10-07 13:42:18","extension":"png","order_by":3,"title":"Figure 3","display":"","copyAsset":false,"role":"figure","size":345936,"visible":true,"origin":"","legend":"\u003cp\u003ePrincipal components analysis using Bray-Curtis Dissimilarity for \u003cstrong\u003e(a)\u003c/strong\u003e all sample types and \u003cstrong\u003e(b)\u003c/strong\u003e lobster larvae and tank water only. Microbial communities associated with early-stage lobster embryos are distinct from those of late-stage embryos and larvae. Late-stage embryos show a wider range of beta-diversity along the axis 1. Microbial communities associated with lobster larvae are most similar to those in tank water (a) but show a higher variance in diversity when analyzed independently (b). Microbial communities in tank water samples are tightly clustered, indicating that they are similar, even among different treatment groups. (Ellipses represent 90% confidence intervals.)\u003c/p\u003e","description":"","filename":"3.png","url":"https://assets-eu.researchsquare.com/files/rs-6831519/v1/e65edc2d18e4db8b2385fa66.png"},{"id":92952915,"identity":"25348e08-d404-4cec-8994-2b76997cc714","added_by":"auto","created_at":"2025-10-07 13:26:18","extension":"png","order_by":4,"title":"Figure 4","display":"","copyAsset":false,"role":"figure","size":199594,"visible":true,"origin":"","legend":"\u003cp\u003eShared core genera. Number of genera shared among lobster samples at 100% prevalence.\u003c/p\u003e","description":"","filename":"4.png","url":"https://assets-eu.researchsquare.com/files/rs-6831519/v1/bd2c3bc4238f8a8ca3197b3f.png"},{"id":102785396,"identity":"07744598-98a0-4aa0-a5ff-1bc652b25571","added_by":"auto","created_at":"2026-02-16 16:06:15","extension":"pdf","order_by":0,"title":"","display":"","copyAsset":false,"role":"manuscript-pdf","size":2006159,"visible":true,"origin":"","legend":"","description":"","filename":"manuscript.pdf","url":"https://assets-eu.researchsquare.com/files/rs-6831519/v1/15738dd4-36c3-485c-8f6e-b4d5e960ec8b.pdf"},{"id":92952909,"identity":"c4e2b109-a368-4a32-9556-7a4993d5570d","added_by":"auto","created_at":"2025-10-07 13:26:17","extension":"docx","order_by":0,"title":"","display":"","copyAsset":false,"role":"supplement","size":21652,"visible":true,"origin":"","legend":"","description":"","filename":"Supp.Table1.docx","url":"https://assets-eu.researchsquare.com/files/rs-6831519/v1/40998e0f135d6008124c999d.docx"},{"id":92953147,"identity":"21152b5a-620f-4f32-9402-320fdb9e07c1","added_by":"auto","created_at":"2025-10-07 13:34:17","extension":"docx","order_by":1,"title":"","display":"","copyAsset":false,"role":"supplement","size":16341,"visible":true,"origin":"","legend":"","description":"","filename":"Supp.Table23.docx","url":"https://assets-eu.researchsquare.com/files/rs-6831519/v1/e9fb1e48cf8e3b13d8b69709.docx"},{"id":92952917,"identity":"9fac1c6e-3d3e-48c6-b6c2-cac47a302d0f","added_by":"auto","created_at":"2025-10-07 13:26:18","extension":"pptx","order_by":2,"title":"","display":"","copyAsset":false,"role":"supplement","size":140840,"visible":true,"origin":"","legend":"","description":"","filename":"SupplementalFigures13.pptx","url":"https://assets-eu.researchsquare.com/files/rs-6831519/v1/25b4c4dcc2e2788735241125.pptx"}],"financialInterests":"No competing interests reported.","formattedTitle":"The bacterial community composition of American lobster (Homarus americanus) embryos and recently hatched larvae held under different temperature and acidification conditions","fulltext":[{"header":"Background","content":"\u003cp\u003eCrustaceans have received modest attention with respect to their bacterial pathogens and symbionts, and even less attention has been directed at their microbiomes. In general, crustacean embryos and larvae are notoriously susceptible to fungal and water mold (oomycete) infections, most notably those caused by species of \u003cem\u003eLagenidium, Saprolegnia, and Haliphthoros\u003c/em\u003e [\u003cspan citationid=\"CR1\" class=\"CitationRef\"\u003e1\u003c/span\u003e]. In the early 1990s, a Gram-negative rod-shaped bacterium isolated from lobster embryos was shown to produce a compound with modest anti-fungal properties. This compound was thought to protect the embryos from fungal infections when present in sufficient concentrations [\u003cspan citationid=\"CR2\" class=\"CitationRef\"\u003e2\u003c/span\u003e, \u003cspan citationid=\"CR3\" class=\"CitationRef\"\u003e3\u003c/span\u003e]. Since then, little progress has been made in characterizing the microbial community associated with lobster eggs. Early research focused on culturable bacteria, yielding only a few isolates without taxonomic identifications. As a result, we have a limited understanding of microbial taxa present during one of the lobster\u0026rsquo;s earliest and most vulnerable life history stages. Given the previous findings of a potential microbial symbiont and the current gap in knowledge, it is important to examine the microbiomes of lobster embryos and larvae using new and advanced techniques.\u003c/p\u003e\u003cp\u003eMicroorganisms associated with a living host form its microbiome, including bacteria, fungi, viruses, archaea, and protists. Microbiomes of marine animals can originate from the surrounding seawater [\u003cspan citationid=\"CR4\" class=\"CitationRef\"\u003e4\u003c/span\u003e] or be transmitted across generations, as seen in embryos of some marine sponges [\u003cspan citationid=\"CR5\" class=\"CitationRef\"\u003e5\u003c/span\u003e, \u003cspan citationid=\"CR6\" class=\"CitationRef\"\u003e6\u003c/span\u003e, \u003cspan citationid=\"CR7\" class=\"CitationRef\"\u003e7\u003c/span\u003e]. Many of the organisms in egg microbiomes are known to be symbiotic, providing metabolic and immune support to the host, protecting the organism from pathogens, or producing secondary metabolic byproducts that support the host\u0026rsquo;s fitness [\u003cspan citationid=\"CR8\" class=\"CitationRef\"\u003e8\u003c/span\u003e]. Conversely, a bloom of pathobionts can slow growth rates, cause pathologies, or compromise host defenses resulting in disease or death to the host [\u003cspan citationid=\"CR9\" class=\"CitationRef\"\u003e9\u003c/span\u003e]. Given the importance of early life history stages, their importance in recruitment to adult populations, and their sensitivity to climate changes, it is surprising that little research has been conducted on the microbiomes of aquatic and marine embryos and their influence on the developing embryo.\u003c/p\u003e\u003cp\u003eAddressing the embryo and larvae microbiome knowledge gap is made more urgent by the impact of climate change. From 1982\u0026ndash;2013, sea surface temperatures in the Gulf of Maine warmed by 0.03\u0026deg;C year\u003csup\u003e\u0026minus;\u0026thinsp;1\u003c/sup\u003e, which is three times faster than the global average rate [\u003cspan citationid=\"CR10\" class=\"CitationRef\"\u003e10\u003c/span\u003e, \u003cspan citationid=\"CR11\" class=\"CitationRef\"\u003e11\u003c/span\u003e]. In general, bacteria thrive in a warmer environment with the potential to double their metabolic rate for every 10\u0026deg;C increase in temperature (i.e., the Q\u003csub\u003e10\u003c/sub\u003e). Some pathogenic species may be able to outpace probiotic species resulting in dysbiosis and disease of their hosts, as observed in American lobsters diagnosed with Epizootic Shell Disease (ESD) [\u003cspan citationid=\"CR12\" class=\"CitationRef\"\u003e12\u003c/span\u003e]. The initial diagnosis of ESD was hampered by the lack of a comparative baseline of the normal shell microbiome. The disease is now known to be a dysbiosis enhanced by the presence of a few chitinolytic species (e.g., \u003cem\u003eAquimarina\u003c/em\u003e sp.) linked to warming water temperature off Southern New England (SNE) [\u003cspan citationid=\"CR13\" class=\"CitationRef\"\u003e13\u003c/span\u003e, \u003cspan citationid=\"CR14\" class=\"CitationRef\"\u003e14\u003c/span\u003e]. The disease has contributed to the decline of the SNE lobster stock [\u003cspan citationid=\"CR15\" class=\"CitationRef\"\u003e15\u003c/span\u003e, \u003cspan citationid=\"CR16\" class=\"CitationRef\"\u003e16\u003c/span\u003e] and has been identified in lobsters in the Gulf of Maine [\u003cspan citationid=\"CR17\" class=\"CitationRef\"\u003e17\u003c/span\u003e]. Due to climate change, it is projected to increase in prevalence at a rapid rate in that region [\u003cspan citationid=\"CR18\" class=\"CitationRef\"\u003e18\u003c/span\u003e]. Although lobster embryos do not appear to be physiologically impacted by ESD [\u003cspan citationid=\"CR19\" class=\"CitationRef\"\u003e19\u003c/span\u003e], it is prudent to establish baseline data on the embryo microbiome prior to the further influences of warmer water, especially as it may provide insight into potential microorganisms important in abnormal clutches. Embryos are arguably the most sensitive and complex stage in the lobster life cycle and are often found dead or dying in the clutches of egg-bearing females. In fact, pathogenic infection is a leading cause of egg mortality in lobsters and other crustaceans [\u003cspan citationid=\"CR20\" class=\"CitationRef\"\u003e20\u003c/span\u003e, \u003cspan citationid=\"CR21\" class=\"CitationRef\"\u003e21\u003c/span\u003e], but the relationship between mortality and the egg microbiome is not yet known.\u003c/p\u003e\u003cp\u003eThis study aims to establish a critical baseline for understanding lobster embryo and larvae microbiomes and their potential resilience or vulnerability to environmental changes. Our objectives were, therefore, a) to characterize the microbial community composition of American lobster embryos, b) to evaluate how embryonic stage, temperature, pH, and origin of the lobsters may influence this community, c) to compare the microbiomes of recently hatched lobster larvae to those of embryos and their surrounding tank environment, and d) to identify the core members of microbiomes in different developmental stages of eggs and the first larval stage. Lobster embryos are extruded (i.e., oviposited) into an external clutch on the female\u0026rsquo;s abdomen and held over a developmental period of 9\u0026ndash;11 months. This extended embryogenesis provides a unique opportunity to investigate microbial community dynamics over time. Recent advances in next generation sequencing now allow us to examine the microbial community of embryos and larvae with unprecedented resolution, quantifying the relative abundance of bacterial taxa at different developmental stages.\u003c/p\u003e"},{"header":"Methods","content":"\u003cdiv id=\"Sec3\" class=\"Section2\"\u003e\u003ch2\u003eLobster Collection \u0026amp; Husbandry\u003c/h2\u003e\u003cp\u003eOvigerous female lobsters were collected by the Massachusetts Division of Marine Fisheries (courtesy of Dr. Tracy Pugh) and the Maine Department of Marine Fisheries (courtesy of Kathleen Reardon, Special License ME 2019-61-03) as part of their ventless trap surveys. Animals were shipped overnight to the Virginia Institute of Marine Science (VIMS) in October 2020, packed in kelp on top of ice bottles. After a brief period of acclimation in a large, chilled, recirculating tank, where they were held together, 24 animals were separated and randomly distributed among four treatment groups (Supplemental Fig.\u0026nbsp;1). Lobster pairs (one from ME waters, one from MA waters) were kept in the same aquaria, but physically separated using plastic bins with a mesh top. Briefly, the control treatment group was kept at 10\u0026ndash;14\u0026deg;C and pH 8.0\u0026ndash;8.1, the warming treatment group was maintained between 14\u0026ndash;18\u0026deg;C with pH 8.0\u0026ndash;8.1, the acidification treatment group at 10\u0026ndash;14\u0026deg;C with a reduced pH 7.5\u0026ndash;7.6, and lastly, the multi-stressor treatment group was kept at 14\u0026ndash;18\u0026deg;C with a pH 7.5\u0026ndash;7.6. Temperatures in all treatment groups were adjusted monthly (hence the range in temperature conditions) to reflect present-day seasonal variation for the southern Gulf of Maine. Each tank contained recirculating seawater (32 psu) that was monitored and controlled autonomously for temperature and pH using a custom system as described in [\u003cspan citationid=\"CR22\" class=\"CitationRef\"\u003e22\u003c/span\u003e]. Water changes (75%) were performed weekly the day after feeding with a combination of filtered seawater from the York River (Gloucester Point, VA) and artificial seawater adjusted to 32 psu made following the manufacturer\u0026rsquo;s protocol (Crystal Sea Marine Mix), as described in detail in [\u003cspan citationid=\"CR22\" class=\"CitationRef\"\u003e22\u003c/span\u003e]. Ovigerous lobsters were fed every 2 weeks with thawed squid during the acclimation period, and then weekly during the treatment period.\u003c/p\u003e\u003c/div\u003e\n\u003ch3\u003eSample Collection and Preparation\u003c/h3\u003e\n\u003cp\u003eAliquots of 10\u0026ndash;20 eggs were aseptically collected from each lobster using forceps in December 2020 (early group) and March 2021 (late group). Stage I larvae were collected within 12 hours of hatching on various dates. Individual, intact eggs and larvae were flash frozen over liquid nitrogen and stored at -80\u0026deg;C. Eggs and larvae were thawed on ice for quantification of protein content. Briefly, they were homogenized with 200uL of MilliEq water and sonicated at at 20% amplitude for 1-second pulses until no tissue was visible (4\u0026ndash;5 pulses). Of the remaining homogenate, 90uL was aseptically removed for determination of protein content, and this procedure is only noted to indicate a freeze-thaw cycle of the homogenized tissue samples. The remaining homogenate was then refrozen and stored at -80\u0026deg;C until DNA extraction for metabarcoding. One egg was used from each ovigerous lobster at each time point. Water samples from each lobster aquarium were collected in 300mL Nalgene bottles near the end point of the experiment in April 2021 and stored at -20\u0026deg; C until thawed for filtration and DNA extraction. To measure bacterial community variation within an egg clutch, two lobsters were selected from which four eggs from each lobster were individually processed as above.\u003c/p\u003e\n\u003ch3\u003eDNA Extraction\u003c/h3\u003e\n\u003cp\u003eThe total DNA was extracted from each remaining tissue homogenate (110 uL, n\u0026thinsp;=\u0026thinsp;32 embryos) using the DNeasy PowerLyzer PowerSoil Kit (Qiagen) following the manufacturer\u0026rsquo;s protocol. DNA quality and concentration were measured using Nanodrop (ThermoFisher Scientific) and Qubit (Life Sciences), respectively. The larvae and tank water samples were processed separately but in the same fashion (n\u0026thinsp;=\u0026thinsp;7 larvae, n\u0026thinsp;=\u0026thinsp;4 tank water). Water samples were thawed on ice and individually filtered with a Sterivex 0.22 um PES syringe filter (Millipore). Each filter was then removed from the housing, aseptically cut into approximately 2 mm\u003csup\u003e2\u003c/sup\u003e pieces, and the DNA extracted from the filter cuttings as above, following the manufacturer\u0026rsquo;s protocol.\u003c/p\u003e\n\u003ch3\u003eAmplicon library preparation\u003c/h3\u003e\n\u003cp\u003eThe extracted DNA from embryos was then diluted to a consistent concentration of 2ng/uL for MiSeq PCR, in which indexed, \u0026ldquo;barcoded\u0026rdquo; MiSeq 515F-Y forward primers and an 806R reverse primer were used to conduct PCR following the Phusion Hi-Fidelity DNA Polymerase procedure. Initial denaturation was carried out at 98\u0026deg;C for 30s, followed by 30 cycles of 10s denaturation at 98\u0026deg;C, 30s of annealing at 55\u0026deg;C and 30s of extension at 72\u0026deg;C. The final extension was carried out at 72\u0026deg;C for 2 minutes. Amplification was confirmed via gel electrophoresis, and the tagged PCR products (n\u0026thinsp;=\u0026thinsp;32 embryos) were pooled and homogenized with 10uL of each product. Aliquots of 30uL of the pooled PCR product were electrophoresed using a 2% agarose, and the resulting band excised from the gel and purified using Wizard SV Gel and PCR Clean-up System (Promega). Amplicon length was assessed using the D1000 ScreenTape system (Agilent). The expected fragment size was approximately 300bp. Thirty-four samples, including one positive and negative control, were sequenced using the Miseq Reagent Kits V2 (250bp reads) on the Illumina MiSeq platform at VIMS. The PCR product from a mock community of known composition (ZymoBIOMICS Microbial Community Standard, Zymo Research Corp.) served as a positive control. Extracted DNA from larvae and tank water samples were amplified and sequenced in the same manner as for the embryos on a subsequent occasion (n\u0026thinsp;=\u0026thinsp;7 larvae, n\u0026thinsp;=\u0026thinsp;4 tank water).\u003c/p\u003e\u003cdiv id=\"Sec7\" class=\"Section2\"\u003e\u003ch2\u003eData Analysis\u003c/h2\u003e\u003cp\u003eThe raw reads were from embryos were trimmed and filtered using the DADA2 analysis package [\u003cspan citationid=\"CR23\" class=\"CitationRef\"\u003e23\u003c/span\u003e] in the R Studio environment [\u003cspan citationid=\"CR24\" class=\"CitationRef\"\u003e24\u003c/span\u003e], as separate analyses for each sequencing run, and merged following chimera removal, as per the DADA2 package manager recommendations. Paired-end reads were trimmed by visually assessing quality scores and by using DADA2\u0026rsquo;s filtering parameters: truncLen\u0026thinsp;=\u0026thinsp;c(200, 160), maxN\u0026thinsp;=\u0026thinsp;0, truncQ\u0026thinsp;=\u0026thinsp;2, rm.phix\u0026thinsp;=\u0026thinsp;TRUE, rm.phix\u0026thinsp;=\u0026thinsp;TRUE, and maxEE\u0026thinsp;=\u0026thinsp;2, and the primers were trimmed using trimLeft\u0026thinsp;=\u0026thinsp;20. Raw reads from tank water and larvae were filtered using the same parameters, except for truncLen\u0026thinsp;=\u0026thinsp;c(150,150). Sequences were dereplicated and sequence variants inferred using the associated error model. Filtered reads were merged and used to build the amplicon sequence variant tables. Denoised full length sequences were trimmed, and chimeras removed. The sequence table from each run was then merged using mergeSequenceTables() in DADA2. Version 132 of the Silva database [\u003cspan citationid=\"CR25\" class=\"CitationRef\"\u003e25\u003c/span\u003e] was used to assign taxonomic affiliations.\u003c/p\u003e\u003cp\u003ePhyloseq [\u003cspan citationid=\"CR26\" class=\"CitationRef\"\u003e26\u003c/span\u003e] and ggplot2 packages [\u003cspan citationid=\"CR27\" class=\"CitationRef\"\u003e27\u003c/span\u003e] in R were used to visualize taxonomic profiles, assess diversity, and perform principal components analysis (PCoA) of the samples. PCoAs were conducted by rarefying samples to the smallest library among the samples and ordinating the distances using Bray-Curtis dissimilarity. All statistical analyses were performed in R, primarily using multivariate community ecology procedures in the base R and vegan [\u003cspan citationid=\"CR28\" class=\"CitationRef\"\u003e28\u003c/span\u003e] packages. A full record of the DADA2 and phyloseq code used, as well as all statistical analyses are included as Additional files one, two and three.\u003c/p\u003e\u003cp\u003eThe core community was defined as the genera which occupied 100% of the lobster samples determined by plotting the number of genera shared for each sample type by their prevalence across samples. The core phyloseq objects were agglomerated by genus, using the function tax_glom(ps, taxrank = \u0026ldquo;Genus\u0026rdquo;). The microbiome package in R [\u003cspan citationid=\"CR29\" class=\"CitationRef\"\u003e29\u003c/span\u003e] was used to calculate genus prevalence using the function: core_members(ps, prevalence\u0026thinsp;=\u0026thinsp;.99). The \u0026lsquo;microbiome\u0026rsquo; package uses prevalence strictly greater than called, thus, a prevalence of 0.99 was used to determine 100% prevalence of genera among the sample types. The code used to analyze the core is included as Additional file four.\u003c/p\u003e\u003c/div\u003e"},{"header":"Results","content":"\u003cdiv id=\"Sec9\" class=\"Section2\"\u003e\u003ch2\u003eEmbryos\u003c/h2\u003e\u003cp\u003eThe average read depth of the embryos was 338,803 with a minimum of 144,986 and a maximum of 452,715. Developing embryos had diverse bacterial communities consisting of 2,317 ASVs associated with 23 embryo samples, after filtering out organisms with chloroplasts and mitochondria. The negative control contained only one read, and the positive control contained the expected taxa in the mock community. Embryos collected in December (early) tended to have lower Perkin\u0026rsquo;s Eye Index (PEI) values than those collected in March (late) (Fig.\u0026nbsp;\u003cspan refid=\"Fig1\" class=\"InternalRef\"\u003e1\u003c/span\u003e). Early embryos had lower Shannon diversity (p\u0026thinsp;\u0026lt;\u0026thinsp;0.05, Kruskal-Wallis test) compared to late embryos (Fig.\u0026nbsp;\u003cspan refid=\"Fig1\" class=\"InternalRef\"\u003e1\u003c/span\u003e). There was no trend observed in early eggs between alpha diversity and PEI, however, alpha diversity increased with PEI in the late samples. The mean Shannon diversity index of early eggs was 2.74 vs. 3.69 for late eggs, with larvae having the highest average index at 4.00. The Simpson and Inverse Simpson index both showed a trend of increasing diversity from early-stage embryos to late-stage embryos to larvae (Supp. table 1). Indices for microbial communities in tank water were slightly above those of early embryos. There were no significant alpha diversity trends noted between the origin of the ovigerous lobster (ME or MA) or among treatment groups (warming or acidification) (\u003cem\u003ep\u003c/em\u003e\u0026thinsp;\u0026gt;\u0026thinsp;0.05, Kruskal-Wallis test). Also of note, there was no difference in these alpha diversity patterns between rarefied and non-rarefied data, although the observed ASVs were generally lower when rarefied for all samples, as expected (Supp. tables 1 \u0026amp; 2).\u003c/p\u003e\u003cp\u003e\u003c/p\u003e\u003cp\u003eTo better visualize the relative abundance data, the dominant microbial taxa associated with lobster embryos were identified at the Order level. Dominant taxa included \u003cem\u003eBacteriodales, Chitinophagales, Microtrichales, Rhizobiales, Betaproteobacteriales\u003c/em\u003e, \u003cem\u003eFlavobacterales\u003c/em\u003e, and \u003cem\u003eVerrucomicrobiales\u003c/em\u003e (Fig.\u0026nbsp;\u003cspan refid=\"Fig2\" class=\"InternalRef\"\u003e2\u003c/span\u003e). Initial analyses indicated that one embryo in the early embryogenesis group, T4L1E2, was highly dissimilar to the others, having a disproportionately high relative abundance of \u003cem\u003eChitinophagaceae\u003c/em\u003e, indicating that this embryo may have been unhealthy. It was removed from statistical analyses as an outlier to prevent disproportionate weighting of that sample.\u003c/p\u003e\u003cp\u003e\u003c/p\u003e\u003cp\u003eMicrobiomes on eggs in both early and late stages of embryogenesis grouped differently but they were the closest associates in community composition when analyzed by PCoA (Fig.\u0026nbsp;\u003cspan refid=\"Fig3\" class=\"InternalRef\"\u003e3\u003c/span\u003ea) (Bray-Curtis Dissimilarity \u003cem\u003ep\u003c/em\u003e\u0026thinsp;\u0026lt;\u0026thinsp;0.05, PERMANOVA). Microbiomes on early-stage embryos showed a more tightly clustered grouping whereas late-stage embryos had a broader range of diversity demonstrated by a wider spread along axis 1 of the PCoA. No other variables such as treatment group or lobster origin showed significant associations in PCoA or via PERMANOVA (\u003cem\u003ep\u003c/em\u003e\u0026thinsp;\u0026gt;\u0026thinsp;0.05, Bray-Curtis Dissimilarity).\u003c/p\u003e\u003cp\u003e\u003c/p\u003e\u003c/div\u003e\n\u003ch3\u003eLarvae\u003c/h3\u003e\n\u003cp\u003eLarvae from seven lobsters were analyzed from five different tanks with representatives from each treatment group (warming, acidification and multi-stressor) and one control. After initial trimming and filtering, 906 ASVs across the seven samples were identified. The average read depth was 33,008 reads with a minimum of 6,535 and a maximum of 87,975. There were several dominant taxa at the order level consisting of \u003cem\u003eFlavobacteriales, Pseudomonadiales, Bacilliales\u003c/em\u003e, and \u003cem\u003eBetaproteobacteriales\u003c/em\u003e. Many taxa enriched in the larvae samples and were not associated with embryos or tank water. Microbiomes of larvae had the highest diversity of all sample types with an average Shannon index of 4.00. Notably, the microbiomes of larvae showed distinctly different community compositions from embryos and the tank water samples, even when the larvae were from different tanks (Figs.\u0026nbsp;\u003cspan refid=\"Fig3\" class=\"InternalRef\"\u003e3\u003c/span\u003ea \u0026amp; \u003cspan refid=\"Fig3\" class=\"InternalRef\"\u003e3\u003c/span\u003eb). In addition, the larvae samples showed no significant trends in community composition in response to the different treatment groups.\u003c/p\u003e\u003cdiv id=\"Sec11\" class=\"Section2\"\u003e\u003ch2\u003eTank water\u003c/h2\u003e\u003cp\u003eTank water was sampled to examine community differences that might arise from the general milieu of the aquaria in which the lobsters and larvae had been held. Tanks one and two were larvae holding tanks, and tanks three and four were aquaria in which lobsters were housed during the experiment. Tank three was a multi stressor group, and tank four was a control group. The water samples were all collected near the endpoint of the experiment, in April or May. The average read depth across the four samples was 141,405 and after quality filtering, denoising and chimera removal, there remained an average of 103,498 reads per sample. After trimming and filtering, 659 ASVs were detected from tank water samples. At the order level, SAR11 clade, \u003cem\u003eFlavobacteriales\u003c/em\u003e, \u003cem\u003eOceanospirallales\u003c/em\u003e and \u003cem\u003eRhodobacterales\u003c/em\u003e dominated the relative abundance of the four samples (Fig.\u0026nbsp;\u003cspan refid=\"Fig2\" class=\"InternalRef\"\u003e2\u003c/span\u003e). The average Shannon index was 2.95.\u003c/p\u003e\u003c/div\u003e\u003cdiv id=\"Sec12\" class=\"Section2\"\u003e\u003ch2\u003eCore community\u003c/h2\u003e\u003cp\u003eThe core community was examined to further understand the stability of taxa present on lobster embryos and larvae and provide a framework for understanding the potential function of the lobster microbiome. We defined the core as the genera present on 100% of the lobster samples (Supplemental Fig.\u0026nbsp;3). By selecting the genera at 100% prevalence, a distinct core community was identified for each life history stage (Fig.\u0026nbsp;\u003cspan refid=\"Fig4\" class=\"InternalRef\"\u003e4\u003c/span\u003e). Early-stage embryos had 24 core members, with 10 genera found only in this life stage: \u003cem\u003eBdellovibrio, Blastopirellula, Bradyrhizobium, Caulobacter, Glaciecola, Loktanella, Paucibacter, Oseudoalteromonas, Truepera, and Vibrionimonas\u003c/em\u003e. Late-stage embryos had 25 core members, 11 of which were exclusive to this life stage: \u003cem\u003eAliisedimentitaria, Altererythrobacter, Aquimarina, Aurantivirga, Blastocatella, Candidatus_Nitrotoga, Cocleimonas, Granulosicococcus, Litoreibacter, Maritalea, and Roseovarius\u003c/em\u003e. Larvae had 11 core members with seven exclusive to this stage: \u003cem\u003eAcinetobacter, Corynebacterium, Lactobacillus, Lawsonella, Prevotella, Staphylococcus, and Streptococcus\u003c/em\u003e. Three genera were shared among all three life history stages; \u003cem\u003eStenotrophomonas, Rubritalea\u003c/em\u003e and \u003cem\u003eDelftia\u003c/em\u003e. Early and late stage embryos shared 10 genera: \u003cem\u003eColwellia, Hellea, Leucothrix, Nitrosomonas, Peredibacter, Pseudrahensia, Pseudomonas, Robiginitomaculum, Sphingorhabdus\u003c/em\u003e, Sva0996_marine_group, and \u003cem\u003eTaibaiella.\u003c/em\u003e\u003c/p\u003e\u003cp\u003e\u003c/p\u003e\u003c/div\u003e\u003cdiv id=\"Sec13\" class=\"Section2\"\u003e\u003ch2\u003eVariation among individual embryos\u003c/h2\u003e\u003cp\u003eTwo lobsters were selected to examine the variation among embryos from the same clutch using four embryos from each lobster, with the resulting microbial compositions analyzed for relative abundance and dissimilarity within and between lobsters. One lobster was from a control group, and the other from a multi-stressor group. All four eggs from each lobster were collected at the same time point. There was 16% variation in beta diversity among the four eggs within an individual, as measured by Bray-Curtis Dissimilarity (Supp. Figure\u0026nbsp;\u003cspan refid=\"Fig2\" class=\"InternalRef\"\u003e2\u003c/span\u003e). In addition, there was no significant difference in diversity between eggs from the same individual when measured by Shannon index (1-sample t-test for outliers, \u003cem\u003ep\u003c/em\u003e\u0026thinsp;\u0026gt;\u0026thinsp;0.05). The limited variation among single embryos from the same clutch confirmed our assumption that the microbial diversity of one embryo is representative of the microbial diversity across an entire clutch was reasonable for this study.\u003c/p\u003e\u003c/div\u003e"},{"header":"Discussion","content":"\u003cp\u003eLobster embryos possess a richly diverse assemblage of bacterial taxa, consisting of 2,561 ASVs. This assemblage changed with embryogenesis, becoming more diverse over developmental time. An increase in Shannon diversity with later developmental stages indicates that the increasing ASV count was not driven by a dominant species type, rather the relative abundances of bacterial taxa were similar among late-stage lobster embryos. Beta-diversity was also driven by life history stage, with early and late embryos harboring distinct communities from larvae and the tank water. All three lobster life history types had uniquely different community structure compared to the surrounding tank water, although communities associated with larvae were more similar to those of the tank water than to other sample types (Figs.\u0026nbsp;\u003cspan refid=\"Fig3\" class=\"InternalRef\"\u003e3\u003c/span\u003ea \u0026amp; \u003cspan refid=\"Fig3\" class=\"InternalRef\"\u003e3\u003c/span\u003eb). This indicates a tight selection process occurring on lobster eggs, where either the host is exerting control through metabolic influences, or the egg coat is acting as a selective surface for microbial colonization and symbiosis. Surprisingly, no associations were observed based on origin, temperature or pH regimes in this experiment which further implies a selection process early, perhaps during initial oviposition, that is independent of or weakly dependent on the conditions of the external environment.\u003c/p\u003e\u003cp\u003eThe consistent increase with development in bacterial alpha and beta-diversity observed with lobster embryos may be attributed to several factors. One possibility is the expansion in embryo size during embryogenesis, which nearly doubles the surface area of the embryo [\u003cspan citationid=\"CR30\" class=\"CitationRef\"\u003e30\u003c/span\u003e], facilitating increased colonization. Another explanation lies in the metabolic waste production and exchange with the external environment throughout embryonic development. The metabolism of lobster embryos slows during the midway point of development, entering a period of diapause during the coldest months where very little, if any, metabolism is occurring [\u003cspan citationid=\"CR31\" class=\"CitationRef\"\u003e31\u003c/span\u003e]. Then, when water temperatures increase, metabolism increases, which pairs with waste production as increased exchange of small molecules with the external environment [\u003cspan citationid=\"CR32\" class=\"CitationRef\"\u003e32\u003c/span\u003e, \u003cspan citationid=\"CR33\" class=\"CitationRef\"\u003e33\u003c/span\u003e]. The embryos in this experiment did not undergo diapause, as the temperature ranges were higher than typical conditions needed for the dormant phase. There was, however, an increase in oxygen consumption observed throughout embryogenesis [\u003cspan citationid=\"CR22\" class=\"CitationRef\"\u003e22\u003c/span\u003e], indicating metabolism of the developing embryo rapidly rises in later stages. The increased metabolism of late-stage lobster embryos might act as a chemical signal or additional substrate, attracting distinctive colonizers later in development and leading to a more diverse community. Developmental rates of the lobster embryos from this experiment did not significantly differ among treatment groups [\u003cspan citationid=\"CR22\" class=\"CitationRef\"\u003e22\u003c/span\u003e]. The implication of this combined work is that not only is the microbiome of lobster embryos resilient to environmental conditions representative of future scenarios of climate change, but the developing larvae within the embryo itself is also seemingly unaffected by these conditions. Finally, members of the early microbiome may act as prey for taxa that colonize older embryos, thereby enriching diversity through shifts in biological relationships within the microbiome itself independent of, or in concert with, the host\u0026rsquo;s increase in waste products.\u003c/p\u003e\u003cp\u003eChange in bacterial community over developmental time is not unique to lobster embryos and was characterized in the hydrothermal vent shrimp, \u003cem\u003eRimicaris exoculata\u003c/em\u003e [\u003cspan citationid=\"CR34\" class=\"CitationRef\"\u003e34\u003c/span\u003e]. However, this system is quite different as mineral deposits on the egg coat of the shrimp also increased with development due to their proximity to hydrothermal vents, providing an alternative colonization substrate. Moreover, these communities are heavily influenced by extreme vent fluid biogeochemistry and maternal aeration in later development. Lobsters in this study were held in artificial seawater with varying water temperature and chemistry indicating the microbiomes on lobster eggs are resilient to environmental conditions within this system, and a change in metabolism from the lobster embryo is likely driving the observed changes in microbiome structure.\u003c/p\u003e\u003cp\u003eMicrobiomes of recently hatched lobster larvae had higher alpha and distinct beta diversity when compared to those of embryos and tank water, perhaps due to the development of the larval gut microbiome upon hatching. No studies have described the gut microbiome in larval, juvenile, or adult American lobsters. One study examining the larval gut microbiome of the European lobster (\u003cem\u003eHomarus Gammarus\u003c/em\u003e) reported a much less diverse assemblage than was found on embryos and larvae in this study, reporting an average Shannon index of around two (exact mean not reported, n\u0026thinsp;=\u0026thinsp;183) [\u003cspan citationid=\"CR35\" class=\"CitationRef\"\u003e35\u003c/span\u003e]. Nonetheless, several of the bacteria identified in the larval samples of this study are known gut colonizers in other species. For example, \u003cem\u003eBacilli\u003c/em\u003e are known gastrointestinal probionts in many organisms, and have been shown to increase gut diversity in the farmed shrimp \u003cem\u003ePenaeus vannamei\u003c/em\u003e when administered with Streptomyces [\u003cspan citationid=\"CR36\" class=\"CitationRef\"\u003e36\u003c/span\u003e]. The European lobster, \u003cem\u003eH. Gammarus\u003c/em\u003e, larvae showed improved growth rates when fed a \u003cem\u003eBacilli\u003c/em\u003e probiotic [\u003cspan citationid=\"CR37\" class=\"CitationRef\"\u003e37\u003c/span\u003e]. Beneficial \u003cem\u003eClostridia\u003c/em\u003e have been found in the gut of farm raised the freshwater prawn, \u003cem\u003eMacrobrachium rosenbergii\u003c/em\u003e [\u003cspan citationid=\"CR38\" class=\"CitationRef\"\u003e38\u003c/span\u003e].\u003c/p\u003e\u003cp\u003eThe microbial community of the tank milieu had lower alpha-diversity compared to those of the lobsters at every stage of development examined and exhibited the most overlap with the larval community. Nonetheless, when microbiomes of larvae and tank water were analyzed independently, beta-diversity was distinct between the sample types indicating the cuticle of the larva is selective for bacteria absent in the seawater, and/or that the gut microbiota is enriched by the nutrients the larvae are filtering from the seawater (the larvae were not fed during this time). These results support previous findings where the diversity of microbes in larvae of the swimming crab (\u003cem\u003ePortunus trituberculatus)\u003c/em\u003e were shown to be unique from the rearing tank water [\u003cspan citationid=\"CR39\" class=\"CitationRef\"\u003e39\u003c/span\u003e] and further supports the selective nature of the decapod shell, be it egg or cuticle.\u003c/p\u003e\u003cp\u003eWe define core members of a microbiome as those simply more prevalent on their hosts than other constituents. They may or may not be beneficial. Embryos and larvae had a diverse core community that was consistent with complex carbon degrading (saprobic) organisms such as those from the families \u003cem\u003eChitinophagaceae and Flavobacteriaceae\u003c/em\u003e, and genus \u003cem\u003eLeucothrix\u003c/em\u003e. \u003cem\u003eLeucothrix\u003c/em\u003e is particularly interesting because it is a known marine fouling organism, often invading crustacean and fish embryos [\u003cspan citationid=\"CR40\" class=\"CitationRef\"\u003e40\u003c/span\u003e], and marine alga [\u003cspan citationid=\"CR41\" class=\"CitationRef\"\u003e41\u003c/span\u003e]. \u003cem\u003eLeucothrix\u003c/em\u003e was identified as a core member in both early and late stage embryos but was found in higher abundance in the early stage. Perhaps the community of early-stage eggs is more susceptible to \u003cem\u003eLeucothrix\u003c/em\u003e infiltration given the lower overall microbial diversity at this stage.\u003c/p\u003e\u003cp\u003eAnother core community member in late embryo samples belonged to the family \u003cem\u003eFlavobacteriaceae\u003c/em\u003e, which comprised 263 ASVs among the samples. Members of this family are common colonizers of marine invertebrates, fish, and alga. Previous genomic analysis in the marine clade of this family revealed several genes encoding polymer-degrading enzymes for complex carbon metabolism, and a high prevalence of genes encoding secondary metabolites such as antioxidants and cytotoxic compounds [\u003cspan citationid=\"CR42\" class=\"CitationRef\"\u003e42\u003c/span\u003e]. The genes found in the marine clade of \u003cem\u003eFlavobacteriaceae\u003c/em\u003e may be important for late-stage lobsters in preventing the overgrowth of fouling organisms susceptible to cytotoxic defenses (i.e. \u003cem\u003eLeucothrix\u003c/em\u003e).\u003c/p\u003e\u003cp\u003eBacteria of the genus \u003cem\u003eAquimarina\u003c/em\u003e are members of the \u003cem\u003eFlavobacteriaceae\u003c/em\u003e family and were associated with only late-stage embryos. The overgrowth of \u003cem\u003eAquimarina spp\u003c/em\u003e. leads to necrotic shell lesions associated with the progression of Epizootic Shell Disease of the juvenile and adult American lobster, particularly in warming waters [\u003cspan citationid=\"CR13\" class=\"CitationRef\"\u003e13\u003c/span\u003e]. The presence of this bacteria on the embryo of lobster, and from adult lobsters without any indication of disease, supports previous research postulating that \u003cem\u003eAquimarina\u003c/em\u003e is typically a non-pathogenic colonizer that can outcompete other species of the carapace microbiome when external conditions are altered (i.e. when water temperature is elevated) [\u003cspan citationid=\"CR14\" class=\"CitationRef\"\u003e14\u003c/span\u003e, \u003cspan citationid=\"CR13\" class=\"CitationRef\"\u003e13\u003c/span\u003e, \u003cspan citationid=\"CR43\" class=\"CitationRef\"\u003e43\u003c/span\u003e].\u003c/p\u003e\u003cp\u003e\u003cem\u003eDelftia, Stenotrophomonas, and Rubritalea\u003c/em\u003e were core members to all life history stages. \u003cem\u003eDelftia\u003c/em\u003e have recently been identified as potential bioremediators, with some marine species able to degrade napthalene [\u003cspan citationid=\"CR44\" class=\"CitationRef\"\u003e44\u003c/span\u003e, \u003cspan citationid=\"CR45\" class=\"CitationRef\"\u003e45\u003c/span\u003e], heavy metals [\u003cspan citationid=\"CR46\" class=\"CitationRef\"\u003e46\u003c/span\u003e], and polycyclic aromatic hydrocarbons (i.e. forever chemicals known as PAHs) [\u003cspan citationid=\"CR47\" class=\"CitationRef\"\u003e47\u003c/span\u003e]. The role \u003cem\u003eDelftia\u003c/em\u003e play on lobster embryos are larvae is likely related to carbon degradation from other sources. \u003cem\u003eDelftia\u003c/em\u003e and \u003cem\u003eStenotrophomonas\u003c/em\u003e have been identified as a core larval member in the hard-shelled mussel (\u003cem\u003eMytilus coruscus\u003c/em\u003e) and was shown to decrease in abundance when experimentally exposed to increased seawater temperature [\u003cspan citationid=\"CR48\" class=\"CitationRef\"\u003e48\u003c/span\u003e]. Stenotrophomonas spp. are known to be important in nutrient cycling, particularly with nitrogen and sulfur [\u003cspan citationid=\"CR49\" class=\"CitationRef\"\u003e49\u003c/span\u003e].\u003c/p\u003e\u003cp\u003e\u003cem\u003eRubritalea\u003c/em\u003e was the final core genus shared between larvae and all embryos, which has previously been identified in ascidia (sea squirt) [\u003cspan citationid=\"CR50\" class=\"CitationRef\"\u003e50\u003c/span\u003e], deep-sea seawater [\u003cspan citationid=\"CR51\" class=\"CitationRef\"\u003e51\u003c/span\u003e], various sponges [\u003cspan citationid=\"CR52\" class=\"CitationRef\"\u003e52\u003c/span\u003e], and a marine gastropod [\u003cspan citationid=\"CR53\" class=\"CitationRef\"\u003e53\u003c/span\u003e]. Not much is known about this genus; however, some species are able to produce secondary metabolites, reduce nitrate, and are red in color due to the carotenoids they contain [\u003cspan citationid=\"CR52\" class=\"CitationRef\"\u003e52\u003c/span\u003e, \u003cspan citationid=\"CR54\" class=\"CitationRef\"\u003e54\u003c/span\u003e]. On sponges and seaweeds, they are thought to play a symbiotic role by scavenging reactive oxygen species or producing antioxidants (55, 56, 57). \u003cem\u003eRubritalea\u003c/em\u003e may play a similar role on developing lobster embryos.\u003c/p\u003e"},{"header":"Conclusion","content":"\u003cp\u003eWe show that lobster eggs maintain a unique and diverse assemblage of bacteria over the course of development. The diversity of this community increases over time, and the eggs share little overlap with the surrounding tank water. Larvae have distinct communities from late-stage embryos, and harbor several unique taxa not detected in the environment around them. In the context of the initial experiment examining warming and acidification, neither the core communities nor the broader microbial communities changed in response to these variables. That is, the bacterial microbiome appeared resilient to changes in temperature and acidification. Our study shows that lobster embryos offer selective and stable microhabitats with which to address questions about the origin of these microbiomes, and the need to examine their longevity and change over the span of embryogenesis in finer detail.\u003c/p\u003e"},{"header":"Abbreviations","content":"\u003cdiv class=\"DefinitionList\"\u003e\u003cdiv class=\"DefinitionListEntry\"\u003e\u003cdiv class=\"Term\"\u003eGoM\u003c/div\u003e\u003cdiv class=\"Description\"\u003e\u003cp\u003eGulf of Maine\u003c/p\u003e\u003c/div\u003e\u003c/div\u003e\u003cdiv class=\"DefinitionListEntry\"\u003e\u003cdiv class=\"Term\"\u003eSNE\u003c/div\u003e\u003cdiv class=\"Description\"\u003e\u003cp\u003eSouthern New England\u003c/p\u003e\u003c/div\u003e\u003c/div\u003e\u003cdiv class=\"DefinitionListEntry\"\u003e\u003cdiv class=\"Term\"\u003eESD\u003c/div\u003e\u003cdiv class=\"Description\"\u003e\u003cp\u003eEpizootic Shell Disease\u003c/p\u003e\u003c/div\u003e\u003c/div\u003e\u003cdiv class=\"DefinitionListEntry\"\u003e\u003cdiv class=\"Term\"\u003ePCoA\u003c/div\u003e\u003cdiv class=\"Description\"\u003e\u003cp\u003ePrincipal Component Analysis\u003c/p\u003e\u003c/div\u003e\u003c/div\u003e\u003cdiv class=\"DefinitionListEntry\"\u003e\u003cdiv class=\"Term\"\u003eCI\u003c/div\u003e\u003cdiv class=\"Description\"\u003e\u003cp\u003eConfidence Interval\u003c/p\u003e\u003c/div\u003e\u003c/div\u003e\u003cdiv class=\"DefinitionListEntry\"\u003e\u003cdiv class=\"Term\"\u003eME\u003c/div\u003e\u003cdiv class=\"Description\"\u003e\u003cp\u003eMaine\u003c/p\u003e\u003c/div\u003e\u003c/div\u003e\u003cdiv class=\"DefinitionListEntry\"\u003e\u003cdiv class=\"Term\"\u003eMA\u003c/div\u003e\u003cdiv class=\"Description\"\u003e\u003cp\u003eMassachusetts\u003c/p\u003e\u003c/div\u003e\u003c/div\u003e\u003cdiv class=\"DefinitionListEntry\"\u003e\u003cdiv class=\"Term\"\u003eDMR\u003c/div\u003e\u003cdiv class=\"Description\"\u003e\u003cp\u003eDepartment of Marine Resources\u003c/p\u003e\u003c/div\u003e\u003c/div\u003e\u003cdiv class=\"DefinitionListEntry\"\u003e\u003cdiv class=\"Term\"\u003eASV\u003c/div\u003e\u003cdiv class=\"Description\"\u003e\u003cp\u003eAmplicon Sequence Variant\u003c/p\u003e\u003c/div\u003e\u003c/div\u003e\u003cdiv class=\"DefinitionListEntry\"\u003e\u003cdiv class=\"Term\"\u003eVIMS\u003c/div\u003e\u003cdiv class=\"Description\"\u003e\u003cp\u003eVirginia Institute of Marine Science\u003c/p\u003e\u003c/div\u003e\u003c/div\u003e\u003cdiv class=\"DefinitionListEntry\"\u003e\u003cdiv class=\"Term\"\u003eDNA\u003c/div\u003e\u003cdiv class=\"Description\"\u003e\u003cp\u003eDeoxyribonucleic Acid\u003c/p\u003e\u003c/div\u003e\u003c/div\u003e\u003cdiv class=\"DefinitionListEntry\"\u003e\u003cdiv class=\"Term\"\u003ePAH\u003c/div\u003e\u003cdiv class=\"Description\"\u003e\u003cp\u003ePolycyclic aromatic hydrocarbon\u003c/p\u003e\u003c/div\u003e\u003c/div\u003e\u003cdiv class=\"DefinitionListEntry\"\u003e\u003cdiv class=\"Term\"\u003ePCR\u003c/div\u003e\u003cdiv class=\"Description\"\u003e\u003cp\u003ePolymerase Chain Reaction\u003c/p\u003e\u003c/div\u003e\u003c/div\u003e\u003cdiv class=\"DefinitionListEntry\"\u003e\u003cdiv class=\"Term\"\u003erRNA\u003c/div\u003e\u003cdiv class=\"Description\"\u003e\u003cp\u003eRibosomal Ribonucleic Acid\u003c/p\u003e\u003c/div\u003e\u003c/div\u003e\u003c/div\u003e"},{"header":"Declarations","content":"\u003cp\u003e\u003cstrong\u003eAvailability of data and materials\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003eThe datasets supporting the conclusions of this article are available at the National Center for Biotechnology Information (NCBI) BioProject repository, accession number PRJNA1227997 (https://www.ncbi.nlm.nih.gov/sra/PRJNA1227997). Data and R code are accessible at https://figshare.com/s/d052cebd1a2ef3f6cb5b. \u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003eCompeting Interests\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003eThe authors declare that they have no competing interests.\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003eFunding\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003eWe thank the following for partial funding of this project: NOAA National Sea Grant’s American Lobster Initiative #NA19OAR4170393 and #NA24OARX417C0578 as well as the Batten School for Coastal and Marine Sciences, Virginia Institute of Marine Science, College of William and Mary.\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003eAuthors Contributions\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003eJSK conceived the study, performed 16S rRNA gene laboratory assays, analyzed the 16S rRNA gene sequencing data, wrote the manuscript, and prepared figures. BS contributed resources and training, and helped with 16S rRNA data analysis. BJ, ARS and ER conceived the climate change study, contributed to animal husbandry, and collected samples. JDS conceived the climate change study, collected samples, wrote the manuscript, and facilitated the project. All authors edited the manuscript and approved the final draft.\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003eAcknowledgements\u003c/strong\u003e:\u003c/p\u003e\n\u003cp\u003eWe would like to thank Drs. J. Goldstein, A. R. Wargo, and K. Reece for their helpful critiques of the manuscript. We are indebted to Kathleen Reardon and her team at the DMR in Maine, and Dr. Tracy Pugh and her team at DMF, Massachusetts for acquiring and shipping lobsters to us. Brett Sweezey and Gabe Thompson helped with lobster maintenance over the course of the experiment. Mara Walters and Lilly Blume contributed by helping with DNA extractions, PCR, helpful discussions, and various other assistance in the lab. This work was funded by the NOAA American Lobster Initiative #NA190AR4170393 to ABR and JDS.\u003c/p\u003e"},{"header":"References","content":"\u003col\u003e\n \u003cli\u003eShields, Jeffrey D., Fran J. Stephens, and Brian Jones. \u0026quot;Pathogens, parasites and other symbionts.\u0026quot;\u003cem\u003eLobsters: biology, management, aquaculture and fisheries\u0026rdquo;\u0026nbsp;\u003c/em\u003e(2006): 146-204.\u003c/li\u003e\n \u003cli\u003eGil-Turnes, M. S., Hay, M. E., \u0026amp; Fenical, W. (1989). Symbiotic marine bacteria chemically defend crustacean embryos from a pathogenic fungus. \u003cem\u003eScience\u003c/em\u003e, \u003cem\u003e246\u003c/em\u003e(4926), 116-118.\u0026nbsp;\u003c/li\u003e\n \u003cli\u003eGil-Turnes, M. S., \u0026amp; Fenical, W. (1992). 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I., Ivanova, A. A., Delegan, Y. A., Streletskii, R. A., Vershinina, D. D., Sokolov, S. L., \u0026amp; Vetrova, A. A. (2023). Characterization and Genomic Analysis of the Naphthalene-Degrading \u003cem\u003eDelftia tsuruhatensis\u003c/em\u003e ULwDis3 Isolated from Seawater. \u003cem\u003eMicroorganisms\u003c/em\u003e, \u003cem\u003e11\u003c/em\u003e(4), 1092.\u003c/li\u003e\n \u003cli\u003eLiu, Y., Tie, B., Li, Y., Lei, M., Wei, X., Liu, X., \u0026amp; Du, H. (2018). Inoculation of soil with cadmium-resistant bacterium \u003cem\u003eDelftia\u003c/em\u003e sp. B9 reduces cadmium accumulation in rice (Oryza sativa L.) grains. \u003cem\u003eEcotoxicology and Environmental Safety\u003c/em\u003e, \u003cem\u003e163\u003c/em\u003e, 223-229.\u003c/li\u003e\n \u003cli\u003eWu, W., Huang, H., Ling, Z., Yu, Z., Jiang, Y., Liu, P., \u0026amp; Li, X. (2016). Genome sequencing reveals mechanisms for heavy metal resistance and polycyclic aromatic hydrocarbon degradation in \u003cem\u003eDelftia lacustris\u003c/em\u003e strain LZ-C. \u003cem\u003eEcotoxicology\u003c/em\u003e, \u003cem\u003e25\u003c/em\u003e, 234-247.\u003c/li\u003e\n \u003cli\u003eZhu, Y. T., Liang, X., Liu, T. T., Power, D. M., Li, Y. F., \u0026amp; Yang, J. L. (2024). The mussel larvae microbiome changes in response to a temperature rise. \u003cem\u003eFrontiers in Marine Science\u003c/em\u003e, \u003cem\u003e11\u003c/em\u003e, 1367608.\u003c/li\u003e\n \u003cli\u003eRyan R.P., Monchy S., Cardinale M., Taghavi S., Crossman L., Avison M.B., Berg G., Van Der Lelie D., Dow J.M. (2009). The versatility and adaptation of bacteria from the genus \u003cem\u003eStenotrophomonas\u003c/em\u003e. \u003cem\u003eNature reviews microbiology\u003c/em\u003e, \u003cem\u003e7\u003c/em\u003e(7), 514-525.\u003c/li\u003e\n \u003cli\u003eYoon, J., Matsuda, S., Adachi, K., Kasai, H., \u0026amp; Yokota, A. (2011). Rubritalea halochordaticola sp. nov., a carotenoid-producing verrucomicrobial species isolated from a marine chordate. \u003cem\u003eInternational journal of systematic and evolutionary microbiology\u003c/em\u003e, \u003cem\u003e61\u003c/em\u003e(7), 1515-1520.\u003c/li\u003e\n \u003cli\u003eSong, J., Lim, Y., Joung, Y., Cho, J. C., \u0026amp; Kogure, K. (2018). Rubritalea profundi sp. nov., isolated from deep-seawater and emended description of the genus \u003cem\u003eRubritalea\u003c/em\u003e in the phylum Verrucomicrobia. \u003cem\u003eInternational journal of systematic and evolutionary microbiology\u003c/em\u003e, \u003cem\u003e68\u003c/em\u003e(4), 1384-1389.\u003c/li\u003e\n \u003cli\u003eScheuermayer, M., Gulder, T. A., Bringmann, G., \u0026amp; Hentschel, U. (2006). \u003cem\u003eRubritalea marina\u003c/em\u003e gen. nov., sp. nov., a marine representative of the phylum \u0026lsquo;Verrucomicrobia\u0026rsquo;, isolated from a sponge (Porifera). \u003cem\u003eInternational journal of systematic and evolutionary microbiology\u003c/em\u003e, \u003cem\u003e56\u003c/em\u003e(9), 2119-2124.\u003c/li\u003e\n \u003cli\u003eYoon, J., Matsuo, Y., Matsuda, S., Adachi, K., Kasai, H., \u0026amp; Yokota, A. (2007). \u003cem\u003eRubritalea spongiae\u003c/em\u003e sp. nov. and\u003cem\u003e\u0026nbsp;Rubritalea tangerina\u003c/em\u003e sp. nov., two carotenoid-and squalene-producing marine bacteria of the family \u003cem\u003eVerrucomicrobiaceae\u003c/em\u003e within the phylum \u0026lsquo;\u003cem\u003eVerrucomicrobia\u0026rsquo;\u003c/em\u003e, isolated from marine animals. \u003cem\u003eInternational journal of systematic and evolutionary microbiology\u003c/em\u003e, \u003cem\u003e57\u003c/em\u003e(10), 2337-2343.\u003c/li\u003e\n \u003cli\u003eKasai, H., A. Katsuta, H. Sekiguchi, S. Matsuda, K. Adachi, K. Shindo, J. Yoon, A. Yokota and Y. Shizuri. 2007. \u003cem\u003eRubritalea squalenifaciens\u003c/em\u003e sp. nov., a squalene-producing marine bacterium belonging to subdivision 1 of the phylum \u0026lsquo;\u003cem\u003eVerrucomicrobia\u003c/em\u003e\u0026rsquo;. \u003cem\u003eInt. J. Syst. Evol. Microbiol.\u003c/em\u003e\u003cstrong\u003e57\u003c/strong\u003e: 1630\u0026ndash;1634.\u003c/li\u003e\n \u003cli\u003eZhuang, Y., Saha, M., Bai, Y., Egan, S., Han, Y., Qiu, Q., ... \u0026amp; Wang, G. (2024). Microbial communities associated with the mature sporophytes and sporelings of the commercially cultivated seaweed \u003cem\u003eSaccharina japonica\u003c/em\u003e in Southern China. \u003cem\u003eJournal of Applied Phycology\u003c/em\u003e, 1-11.\u003c/li\u003e\n \u003cli\u003eShindo, K., Asagi, E., Sano, A., Hotta, E., Minemura, N., Mikami, K., Tamesada E., \u0026amp; Maoka, T. (2008). Diapolycopenedioic acid xylosyl esters A, B, and C, novel antioxidative glyco-C30-carotenoic acids produced by a new marine bacterium \u003cem\u003eRubritalea squalenifaciens.\u003c/em\u003e\u003cem\u003eThe Journal of Antibiotics\u003c/em\u003e, \u003cem\u003e61\u003c/em\u003e(3), 185-191.\u003c/li\u003e\n \u003cli\u003eYoon, J., Matsuo, Y., Matsuda, S., Adachi, K., Kasai, H., \u0026amp; Yokota, A. (2008). \u003cem\u003eRubritalea sabuli sp\u003c/em\u003e. nov., a carotenoid-and squalene-producing member of the family \u003cem\u003eVerrucomicrobiaceae\u003c/em\u003e, isolated from marine sediment. \u003cem\u003eInternational journal of systematic and evolutionary microbiology\u003c/em\u003e, \u003cem\u003e58\u003c/em\u003e(4), 992-997.\u003c/li\u003e\n\u003c/ol\u003e"}],"fulltextSource":"","fullText":"","funders":[],"hasAdminPriorityOnWorkflow":false,"hasManuscriptDocX":true,"hasOptedInToPreprint":true,"hasPassedJournalQc":"","hasAnyPriority":false,"hideJournal":false,"highlight":"","institution":"","isAcceptedByJournal":true,"isAuthorSuppliedPdf":false,"isDeskRejected":"","isHiddenFromSearch":false,"isInQc":false,"isInWorkflow":false,"isPdf":false,"isPdfUpToDate":true,"isWithdrawnOrRetracted":false,"journal":{"display":true,"email":"[email protected]","identity":"scientific-reports","isNatureJournal":false,"hasQc":true,"allowDirectSubmit":false,"externalIdentity":"scirep","sideBox":"Learn more about [Scientific Reports](http://www.nature.com/srep/)","snPcode":"","submissionUrl":"","title":"Scientific Reports","twitterHandle":"","acdcEnabled":true,"dfaEnabled":true,"editorialSystem":"stoa","reportingPortfolio":"Scientific Reports","inReviewEnabled":true,"inReviewRevisionsEnabled":true},"keywords":"microbiome, climate change, Crustacea, metabarcoding","lastPublishedDoi":"10.21203/rs.3.rs-6831519/v1","lastPublishedDoiUrl":"https://doi.org/10.21203/rs.3.rs-6831519/v1","license":{"name":"CC BY 4.0","url":"https://creativecommons.org/licenses/by/4.0/"},"manuscriptAbstract":"\u003cp\u003ePrevious research investigating the microbial community of American lobster embryos has long led researchers to believe this habitat comprised only a select few bacterial taxa. However, using 16S rRNA gene sequencing, we show this community to be more diverse than previously thought. We investigated how the bacterial communities of American lobster embryos and larvae change over embryogenesis and hatching in response to two environmental variables. Ovigerous female lobsters caught from Maine and Massachusetts were held under varying temperature and pH regimes that approximated observed and predicted warming and ocean acidification conditions in the Gulf of Maine (GoM) and Southern New England (SNE). The bacterial microbiome associated with the lobster embryos was quantified from two-time points during the experiment, and larvae were collected within 12 hours of hatching. Alpha diversity increased with each life history stage, and embryo and larvae microbiomes shared little community overlap with that in the surrounding tank water. Neither environmental conditions nor lobster origin significantly altered bacterial communities, with life history stage driving alpha and beta diversity. Embryos and larvae shared three core bacterial members identified as members of the genera \u003cem\u003eRubritalea, Delftia\u003c/em\u003e, and \u003cem\u003eStenotrophomonas\u003c/em\u003e. American lobster embryos and larvae appear to have a highly selective microhabitat for bacteria that is not altered by environmental conditions. This leads us to wonder what role the microbiome may have on a developing lobster, and where the microbiome is originating if not from the surrounding seawater.\u003c/p\u003e","manuscriptTitle":"The bacterial community composition of American lobster (Homarus americanus) embryos and recently hatched larvae held under different temperature and acidification conditions","msid":"","msnumber":"","nonDraftVersions":[{"code":1,"date":"2025-10-07 13:26:13","doi":"10.21203/rs.3.rs-6831519/v1","editorialEvents":[{"type":"communityComments","content":0},{"type":"decision","content":"Revision requested","date":"2025-10-29T21:24:30+00:00","index":"","fulltext":""},{"type":"editorInvitedReview","content":"","date":"2025-10-29T17:59:52+00:00","index":"hide","fulltext":""},{"type":"editorInvitedReview","content":"","date":"2025-10-27T15:06:16+00:00","index":"hide","fulltext":""},{"type":"reviewerAgreed","content":"216477257345605774195583719763367850938","date":"2025-10-22T13:43:21+00:00","index":"hide","fulltext":""},{"type":"editorInvitedReview","content":"","date":"2025-10-18T10:05:55+00:00","index":"hide","fulltext":""},{"type":"reviewerAgreed","content":"310298764128472860081219959748020300304","date":"2025-10-17T16:12:12+00:00","index":"hide","fulltext":""},{"type":"editorInvitedReview","content":"","date":"2025-10-16T17:29:09+00:00","index":"hide","fulltext":""},{"type":"reviewerAgreed","content":"130707539569403825654364181245851784052","date":"2025-09-27T09:30:13+00:00","index":"hide","fulltext":""},{"type":"reviewerAgreed","content":"333092552607500439925016279572204975708","date":"2025-09-27T09:08:02+00:00","index":"hide","fulltext":""},{"type":"reviewerAgreed","content":"188272798394432268605453979189719065215","date":"2025-09-25T09:39:14+00:00","index":"hide","fulltext":""},{"type":"reviewersInvited","content":"","date":"2025-09-24T18:47:15+00:00","index":"","fulltext":""},{"type":"editorInvited","content":"","date":"2025-06-10T13:23:48+00:00","index":"","fulltext":""},{"type":"editorAssigned","content":"","date":"2025-06-08T04:05:37+00:00","index":"","fulltext":""},{"type":"checksComplete","content":"","date":"2025-06-06T12:14:51+00:00","index":"","fulltext":""},{"type":"submitted","content":"Scientific Reports","date":"2025-06-05T18:35:15+00:00","index":"","fulltext":""}],"status":"published","journal":{"display":true,"email":"[email protected]","identity":"scientific-reports","isNatureJournal":false,"hasQc":true,"allowDirectSubmit":false,"externalIdentity":"scirep","sideBox":"Learn more about [Scientific Reports](http://www.nature.com/srep/)","snPcode":"","submissionUrl":"","title":"Scientific Reports","twitterHandle":"","acdcEnabled":true,"dfaEnabled":true,"editorialSystem":"stoa","reportingPortfolio":"Scientific Reports","inReviewEnabled":true,"inReviewRevisionsEnabled":true}}],"origin":"","ownerIdentity":"9f133668-c83a-42c7-9280-8023f069cd83","owner":[],"postedDate":"October 7th, 2025","published":true,"recentEditorialEvents":[],"rejectedJournal":[],"revision":"","amendment":"","status":"published-in-journal","subjectAreas":[{"id":55774498,"name":"Biological sciences/Microbiology"},{"id":55774499,"name":"Biological sciences/Microbiology/Communities"},{"id":55774500,"name":"Biological sciences/Ecology/Microbial ecology"}],"tags":[],"updatedAt":"2026-02-16T16:03:23+00:00","versionOfRecord":{"articleIdentity":"rs-6831519","link":"https://doi.org/10.1038/s41598-026-35387-8","journal":{"identity":"scientific-reports","isVorOnly":false,"title":"Scientific Reports"},"publishedOn":"2026-02-14 15:57:16","publishedOnDateReadable":"February 14th, 2026"},"versionCreatedAt":"2025-10-07 13:26:13","video":"","vorDoi":"10.1038/s41598-026-35387-8","vorDoiUrl":"https://doi.org/10.1038/s41598-026-35387-8","workflowStages":[]},"version":"v1","identity":"rs-6831519","journalConfig":"researchsquare"},"__N_SSP":true},"page":"/article/[identity]/[[...version]]","query":{"redirect":"/article/rs-6831519","identity":"rs-6831519","version":["v1"]},"buildId":"8U1c8b4HqxoKbykW_rLl7","isFallback":false,"isExperimentalCompile":false,"dynamicIds":[84888],"gssp":true,"scriptLoader":[]}