Gene expression changes associated with the developmental plasticity of sea urchin larvae in response to food availability.
Developmental plasticity is the environmental elicitation of alternate phenotypes from a single genotype, which facilitates the optimization of a match between phenotype and environment (Dewitt et al., 1998; Agrawal, 2001; Gilbert and Epel, 2009). Marine larvae can experience significant fluctuations in temperature, pressure, salinity, nutrition, and predation. Of these factors, food availability is particularly influential in the development of marine invertebrate larvae (Strathmann et al., 1992; Byrne et at, 2008; Soars et al., 2009).
Planktotrophic echinoid larvae (echinoplutei) are provided with limited maternal investment and rely heavily on exogenous resources (e.g., phytoplankton) for growth to competency for metamorphosis. During pelagic transport, larvae often experience growth-limiting conditions owing to scarcity of food (Conover, 1968; Olson and Olson, 1989). When echinoplutei encounter such conditions, development of the juvenile rudiment is suppressed, while the post-oral arms lengthen and the stomach shrinks (Fenaux et al., 1988; Strathmann et al., 1992; Miner, 2005). This response was recently shown to be mediated by a decrease in dopamine signaling (Adams et al., 2011). Furthermore, it has been shown that thyroid-hormone-like compounds obtained from the microalgae upon which echinoplutei feed promote rudiment development and metamorphosis (Heyland and Hodin, 2004; Heyland et al., 2004). In the face of food scarcity, sea urchin larvae may make use of their ability to take up free amino acids for subsistence (Manahan et al., 1983).
Although the food-responsive developmental plasticity of sea urchin larvae has been examined morphometrically (Boidron-Metairon, 1988; Strathmann et al, 1992; Heyland and Hodin, 2004; Miner, 2005; McAlister, 2008) and molecularly (Adams et al, 2011), genome-wide changes in gene expression associated with this phenomenon have not been explored. To begin filling this void, we used next-generation, high-throughput RNA sequencing (RNA-Seq) as well as quantitative reverse transcription and polymerase chain reaction (qRT-PCR) to investigate the gene expression changes that occur in larvae of the sea urchin Strongylocentrotus droebachiensis O. F. Muller, 1776 in response to different feeding regimes. Additionally, we tested the hypothesis that TOR and FoxO, key regulatory molecules known to control nutrient signaling, responsiveness to energy stress, plasticity, and aging in a variety of animal models, similarly contribute to the food-responsive developmental plasticity of echinoplutei.
Materials and Methods
Procurement of gametes, larval culture, and preparation of RNA
Gametes of Strongylocentrotus droebachiensis from a parental mixture were obtained and fertilized at the University of Maine's Center for Cooperative Aquaculture Research (CCAR; Franklin, ME), using standard methods. Embryos were cultured at CCAR in 18-1 acrylic plastic hatching conical vessels at 50 eggs [ml.sup.-1] until feeding commenced, at which time the embryos were collected by filtration through a 105-[micro]m screen and transferred into fluorescently backlit, flow-through 230-1 fiberglass cylinders at 4 larvae [ml.sup.-1]. Some of the larvae were transferred to the MDI Biological Laboratory (MDIBL) and cultured at 4 larvae [ml.sup.-1] in 1 liter of filtered seawater in magnetic stir jars at 8 [degrees]C.
Larvae were fed a 50:50 mixture of Dunaliella tertiolecta and Rhodomonas salina. For the RNA-Seq experiment, ad libitum feeding (>20,000 cells [larva.sup.-1] [ml.sup.-1]) was carried out at CCAR, and diet-restricted feeding (200 cells [larva.sup.-1] [ml.sup.-1]) was carried out at MDIBL. For RNA-Seq, samples of larvae fed ad libitum were flash-frozen at 2 days post-fertilization (dpf) (gastrula), 6 dpf (pre-feeding pluteus), 13 dpf (6-arm pluteus), 22 dpf (8-armed pluteus with rudiment), and 27 dpf (8-arm pluteus competent for metamorphosis); samples of diet-restricted larvae were flash-frozen at 2 dpf (gastrula), 7 dpf (pre-feeding pluteus), and 38 dpf (6-armed pluteus). RNA was then extracted from each sample using RNaqueous-Midi total RNA isolation kit (Ambion) with lithium chloride precipitation, and quality-tested using a 2100 Bioanalyzer system (Aglient Technologies). Isolated RNA was sent to the Sick Kids Centre for Applied Genomics at the University of Toronto where Illumina mRNA-Seq libraries were prepared for each sample and sequenced on an Illumina GAIIx using manufacturer's protocols.
In a second experiment, using the procedure described above, 6-armed (21 days post-fertilization) larvae at 2 larvae [ml.sup.-1] were fed a 50:50 D. tertiolecta and R. salina mixture every 3 days at 50, 500, and 5000 cells [larva.sup.-1] [ml.sup.-1]. An additional culture was fed 5000 cells [larva.sup.-1] [ml.sup.-1] in the presence of 100 nmol 1[1.sup.-1] rapamycin, a concentration determined by a survey of the literature to be used routinely to inhibit TOR activity in mammalian cells (e.g., Almilaji et al., 2012), and which we showed in preliminary experiments to inhibit rudiment development while having no adverse effect on larval somatic growth or survival. Rapamycin was added to the culture with each water change, which was done prior to each feeding. Once a week, 100 larvae were removed from each culture and flash-frozen. RNA was purified from the frozen aliquots using the RNeasy plus mini kit (Qiagen), and quantified using a Nanodrop ND-1000 spectrophotometer.
Metabolic rates for the larvae used in the RNA-Seq experiment were estimated by measuring the rate of decline in dissolved oxygen in a sealed, stirred, and temperature-controlled chamber containing 1000 larvae. The larvae were washed free of external microalgae by centrifugation prior to being introduced into the chamber, although the stomachs of the well-fed larvae remained full of microalgae. Dissolved oxygen levels were measured using an Ocean Optics fluorescence-based oxygen sensor (FOXY) and multifrequency phase fluorometer.
mRNA-Seq data analysis
Sequence reads were analyzed using CLCBio Genomics Workbench (ver. 4.0) using the following workflow. First, the reads were trimmed by quality after examining the sequence quality and composition of the reads. Next, trimmed reads were mapped to the Strongylocentrotus purpuratus NCBI ver. 2.1 genome assembly, and expression levels for each gene were reported in reads per kilobase of exon model (RPKM) units. Mapped reads were at least 80% identical to the S. purpuratus assembly over at least 50% of their trimmed length. Reads were mapped to the S. purpuratus genome assembly in lieu of de novo transcriptome assembly because S. droebachiensis is a close relative of S. purpuratus (Lee, 2003), so a majority of reads from the former would be expected to map to annotated genes of the latter (borne out by the results, see below and Table S1, http://www.biolbull.org/content/supplemental), permitting differential expression analysis. Enriched Gene Ontology annotations among differentially expressed genes were calculated using the R/topGO (Alexa and Rahnenfuhrer, 2010) (ver. 2.16.0) package in the R (ver. 3.1.1) statistical computing environment, taken from Ensembl Genomes (Kersey et al., 2014) (ver. 23) annotation of S. purpuratus genes obtained using BioMart (Kinsella et al., 2011). For the enrichment analysis, all expressed genes were used as the background.
Quantitative reverse transcription and polymerase chain reaction (qRT-PCR)
RNA was reverse-transcribed to cDNA using Superscript III (Life Technologies) and random hexamer primers, and expression levels for genes of interest were measured in triplicate by qPCR with Perfecta SYBR Green (Quanta Biosciences). Relative expression levels were calculated using the delta-delta Ct method, using 3-UTR sequences from hprt (hypoxanthine-guanine phosphoribosyltransferase; expression of which was not affected by the experimental treatments), as a reference for normalization. Primer sequences were obtained from sequence contigs retrieved by BLAST queries of the S. droebachiensis transcriptome for each gene of interest (Table S4, http://www.biolbull.org/content/supplemental), using the program Primer3Plus. The sequences of the primers are provided in Table 1.
Morphometrics and assessment of competency for metamorphosis
After 4 weeks of differential feeding, 10 larvae arbitrarily selected from each diet were imaged on a Zeiss Axiovert 25 microscope equipped with a Zeiss AxoiCam MRm. Rudiment area was measured using NIH ImageJ software, ver. 1.9.2 (http://imagej.nih.gov/ij/). To determine competency for metamorposis, 50 larvae per diet were removed from the culture and allowed to settle in glass dishes for 72 h and monitored for metamorphosis. Post-metamorphic test size was measured after imaging of the metamorphosed juveniles as described above.
RNA-Seq analysis of larval gene expression under different feeding regimes
To assess the effect of different feeding regimes on the Strongylocentrotus droebachiensis larval transcriptome, we performed mRNA-Seq on total RNA extracted from larvae fed either ad libitum or a restricted diet (see Materials and Methods). Samples of the ad libitum (AL) culture were taken at 2, 6, 13, 22, and 27 days post-fertilization (dpf); larvae were at the 6-arm stage at 13 dpf, and achieved metamorphosis by ~28 dpf. Samples of the diet-restricted (DR) culture were taken at 2, 7, and 38 dpf; larvae were at the 6-arm stage at 38 dpf (Fig. 1). Measurements of oxygen consumption rates of the sampled larvae suggested that the metabolic rate of DR larvae at 38 dpf was about half that of equivalently staged (6-arm) AL larvae at 13 dpf (data not shown), although whether this was due to intrinsic metabolic differences or simply to the fact that the stomachs of the AL larvae were full of microalgae that may still have been respiring cannot be ascertained.
An average of 44.4 million 76-bp reads were generated per sample, and an average of 57.8% of trimmed reads per sample were mapped to the Strongylocentrotus purpuratus genome assembly (Sodergren et al., 2006) (Table S1, http://www.biolbull.org/content/supplemental), with an average of 84.7% of mapped reads per sample mapping to annotated exons. Comparing expression levels between 38-dpf DR samples and 13-dpf AL samples relative to their respective 2-dpf samples identified 2349 genes that were up-regulated and 1966 genes that were down-regulated in response to starvation (Table S2, http://www.biolbull.org/content/supplemental). These genes were differentially expressed by at least 2-fold and had more than five reads per kilobase of exon gene model (RPKM) in at least one sample. The functional context of the up- and down-regulated genes was analyzed by examining enriched Gene Ontology (Ashburner et al., 2000) Biological Process (BP) terms in REVIGO, October 2014 (Supek et al., 2011) (Table S3, http://www.biolbull.org/content/supplemental). Lipid transport and its descendent BP terms constituted the largest category of enriched terms for up-regulated genes (Fig. S1 A, http://www.biolbull.org/content/supplemental). The three largest categories of enriched BP terms for down-regulated genes were DNA conformation change (including gene expression and DNA and RNA metabolism), cellular component biogenesis, and respiratory electron transport chain (Fig. S1B, http://www.biolbull.org/content/supplemental). In general, these changes suggest that starved larvae down-regulate growth-related processes such as protein synthesis and organelle biogenesis that entail a high metabolic demand, while up-regulating processes involved in lipid mobilization, membrane biology, and responsiveness to environmental stimuli (e.g., neurotransmitter transport, ion transport, and signal-transduction-related processes).
The three transcription factors found to be most highly over-expressed in underfed, 38-day-old 6-arm plutei compared to their well-fed 13-day-old counterparts were CREB, Elk, and FoxJ1 (Fig. 2). Genetic variability between the two cultures is unlikely to account for this differential expression, as none of these factors was differentially expressed between the respective 2-day (gastrula stage) samples. Furthermore, after the onset of feeding, Elk and FoxJ1 in the well-fed larvae showed constant per-larva expression throughout development to metamorphosis, suggesting that the high level of overexpression observed in the 38-day underfed larvae was due to the different feeding regime rather than to developmental or chronological time. Expression of CREB increased with development past the 6-arm stage in the well-fed larvae, which correlated temporally with development of the rudiment. However, the 38-day underfed larvae lacked rudiments, suggesting that overexpression of CREB in the latter occurred specifically in response to food scarcity. This fits with what is known from studies of mammals and flies, wherein CREB functions in the regulation of energy balance and mobilization of energy stores in response to fasting or starvation (Iijima et al., 2009; Oh et al., 2013).
Validation of RNA-Seq results by quantitative RT-PCR
To validate and extend the RNA-Seq findings, we performed an experiment in which a culture of 3-week-old, 6-arm larvae developed from a single batch of embryos was divided evenly into four cultures, three of which were respectively fed with 50, 500, and 5000 cells of microalgae [larva.sup.-1] [ml.sup.-1] biweekly; the fourth was fed 5000 cells [larva.sup.-1] [ml.sup.-1] biweekly in the presence of rapamycin, an inhibitor of the nutrient-sensing kinase TOR. After 3 weeks of these treatments, larvae in each culture had developed to the 8-armed stage. However, by this time most larvae on the low-food diet ("starved" larvae) as well as those treated with rapamycin either lacked or had very small rudiments. In contrast, those fed intermediate and high-food diets had well-developed rudiments (Fig. 3 and Fig. 4A), and by 4 weeks had achieved competency for metamorphosis (Fig. 4B). Post-metamorphic test diameter of larvae on medium and high-food diets did not significantly differ, but rapamycin-treated larvae that underwent metamorphosis displayed nearly 2-fold reduction in test diameter (Fig. 4C).
Quantitative RT-PCR was used to measure the expression of several of the genes found to be differentially expressed by RNA-Seq: the transcription factors Elk, FoxJ1, CREB (Fig. 2), as well as NFKB (RNA-Seq indicating ~ 1.9-fold higher expression in starved larvae); the neural excitatory amino acid transporter EAAT (~4-fold higher expression in starved larvae); the translation inhibitor 4E-BP (~2-fold higher expression in starved larvae); the ribosomal biosynthesis factor nucleolin (Nuc, ~2-fold lower expression in starved larvae); and the mitochondria-associated proteins ATP synthase-coupling factor 6 (ATPSyn, ~2.6-fold lower expression in starved larvae), Cytochrome c (CytC,--3.8-fold lower expression in starved larvae), and HSPE1 (--3.4-fold lower expression in starved larvae). In comparison to both intermediate and well-fed larvae, Elk (P < 0.008), CREB (P < 0.0001), FoxJ1 (P < 0.0005), NFKB (P < 0.005), EAAT (P < 0.0005), 4E-BP (P < 0.0005), and ATPSyn (P < 0.0003) expression levels were significantly elevated in starved larvae, each except for ATPSyn recapitulating what was observed in the RNA-Seq experiment (Fig. 5). Expression levels of those genes also significantly increased (P [less than or equal to] 0.015) in the well-fed larvae treated with rapamycin, phenocopying the situation in starved larvae (Fig. 5). Starved and rapamycin-treated larvae displayed no differential expression of ATPSyn (P > 0.35), EAAT (P > 0.07), or Elk (P > 0.30). In starved larvae, CytC (P < 0.001), Nuc (P < 0.005), and HSPE1 (P < 0.00001) expression was significantly decreased, but only HSPE1 (P < 0.0003) expression significantly decreased in rapamycin-treated larvae, while CytC (P > 0.15) remained constant, and Nuc (P < 0.00001) significantly increased.
It should be noted that the experimental context for the comparisons made by qRT-PCR was somewhat different than that made by RNA-Seq, in that the latter compared larvae of the same morphological stage (6-arm) but different ages (well-fed larvae at 13 dpf vs. underfed larvae at 38 dpf), whereas the former compared larvae of the same age (6 weeks) but different morphological stages (well-fed 8-arm larvae with rudiments vs. underfed 8-arm larvae without rudiments). This raises the possibility that the differences in gene expression observed by qRT-PCR might simply be due to spatially differential gene expression between the larval soma and the juvenile rudiment. However, this possibility is ruled out by both the RNA-Seq data (Fig. 2) and the time-course qRT-PCR data presented below (Fig. 6), showing that regulatory genes that are highly overexpressed in response to starvation do not display any decrease in expression in well-fed larvae between the 6-arm stage and the 8-arm stage with a fully developed rudiment.
Time-course of FoxO and 4E-BP expression
In Drosophila the translational inhibitor 4E-BP has been shown to be upregulated in response to nutritional stress, providing a metabolic "brake" to control fat metabolism (Teleman et al., 2005). 4E-BP is post-translationally inhibited by TOR kinase and transcriptionally activated by FoxO, which mediates plasticity in response to decreased growth factor (e.g., insulin) signaling (Puig et al., 2003; Tang et al, 2011). In addition to being post-translationally regulated, FoxO is known to positively regulate its own transcription in some contexts (Essaghir et al, 2009). To examine the effect of different feeding regimes on FoxO expression and activity, we employed qRT-PCR to measure expression of both FoxO and 4E-BP over the entire course of the experiment. Larvae exposed to different feeding regimes displayed dose-dependent differential expression of both FoxO and 4E-BP, with higher expression levels in starved larvae (Fig. 6). Well-fed larvae treated with rapamycin did not follow this trend; their expression profiles more closely resembled those in diet-restricted larvae, albeit with a temporal lag (Fig. 6). After 4 weeks, larvae cultured with rapamycin showed the largest increase in FoxO expression (~10 fold; P < 0.0001), and those cultured on a low-food diet also showed a greater (~7 fold; P < 0.0001) increase in FoxO than those cultured on intermediate- and high-food diets (~5 fold; P < 0.0001) (Fig. 6). 4E-BP displayed similar trends in expression, except that expression in starved larvae was greater than in rapamycin-treated larvae at the end of 4 weeks (Fig. 6). After 4 weeks, 4E-BP expression increased most in starved larvae (~18 fold; P < 0.0001), followed by rapamycin-treated (~10 fold; P < 0.0001), intermediate- (~7 fold; P < 0.0001), and high-food (~5 fold; P < 0.0001) diets.
Plasticity in response to food availability serves to produce a better fit between phenotype and environment, through changes in individual chemistry, physiology, development, morphology, or behavior (Dewitt et al., 1998; Agrawal, 2001; Gilbert and Epel, 2009). To increase fitness and food clearance rate during periods of low food availability, sea urchin larvae elongate post-oral arms and reabsorb stomach tissues; in contrast, when food is abundant, post-oral arms remain short while the stomach lengthens (Strathmann et al, 1992; Miner, 2005; Adams et al., 2011). This adaptation appears to be widely conserved among echinoids (Strathmann et al., 1992; Heyland and Hodin, 2004; Miner, 2005; Reitzel and Heyland, 2007; Byrne et al, 2008; Adams et al., 2011), although subject to evolutionary variation (McAlister, 2008).
We found that starved 38-day echinoplutei of the sea urchin Strongylocentrotus droebachiensis down-regulate genes associated with growth and mitochondrial activity, consistent with preliminary respirometry measurements suggesting that they have a lower metabolic rate than that of well-fed 13-day larvae of an equivalent morphological stage. Concomitantly, the starved larvae up-regulate genes that are known in other animals to regulate energy homeostasis (e.g., CREB, 4E-BP, FoxO), environmental sensing and neuroplasticity (e.g., CREB, Elk, EAAT), ciliagenesis (e.g., FoxJ1), and stress-resistance (e.g., FoxO, NFKB) (Amara and Fontana, 2002; Li and Stark, 2002; Chloe and Wang, 2002; Carter and Brunet, 2007; Iijima et al., 2009; Besnard et al., 2011; Hay, 2011; Oh et al., 2013; Choksi et al., 2014; Webb and Brunet, 2014). This suggests that during periods of nutritional hardship, echinoplutei enhance their sensitivity to environmental conditions and capacity to defend against stress.
Studies using Caenorhabditis elegans and Drosophila have shown that diet restriction increases activity of FoxO and its target 4E-BP while decreasing TOR activity, which increases lifespan; these effects are reversed when food is abundant (Teleman et al, 2005; Hay, 2011; Webb and Brunet, 2014). Our data suggest that these regulatory relationships are conserved in sea urchin larvae and control developmental transit toward metamorphosis. In addition to correlating with nutrient supply and decreased FoxO activity, rudiment growth and/or development is inhibited by rapamycin, suggesting that it requires TOR activity. It is interesting to consider this finding in light of the emerging consensus that the gene encoding TOR, a pro-growth kinase that is essential for early development, is a key pro-aging gene in adult animals, epitomizing the criteria for the antagonistic pleiotropy theory of aging proposed by George Williams in 1957 (Williams, 1957; Blagosklonny, 2010). Our findings are consistent with Williams' prediction that arresting development prior to reproductive maturation would prevent senescence: inhibiting TOR, a kinase that promotes cell proliferation, growth, and protein synthesis (Wullschleger et al, 2006), suppresses development of the adult rudiment (by as-yet-unknown mechanisms) while allowing continued larval existence, possibly indefinitely.
In the wild, suspension of rudiment development in response to food scarcity could allow echinoplutei to serve as "life rafts" for dispersive colonization, resuming development when food becomes available. In support of this idea we have found that after 7 months on a restricted diet precluding rudiment development, S. purpuratus larvae remain competent to develop through metamorphosis when provided sufficient food (Davis and Coffman, 2011). We have been able to maintain diet-restricted larvae for up to a year in the laboratory (JAC, unpubl. results). The maximum lifespan of echinoplutei remains an open question, as does, given the ability of these larvae to undergo cloning (Eaves and Palmer, 2003; Vaughn and Strathmann, 2008), the possibility that they afford echinoids the potential for metagenesis (Mortensen, 1921).
Nevertheless it is clear that echinoid larvae, as well as planktonic larvae of other marine invertebrates, can remain planktonic long enough to cross oceans and maintain connectivity between populations (Scheltema, 1977; Strathmann, 1978; Todd et al., 1998; Behnam et al., 2012), suggesting that starvation-induced suspension of development is an effective mechanism for increasing gene flow across meta-populations and gyres. Teleplanic larvae of echinoderms, molluscs, polychaetes, and crustaceans (Scheltema, 1971; Strathmann, 1978; Shanks, 2009) may remain in the open sea for more than 300 days, and veligers of the snail Fusitriton oregonensis have been maintained in the laboratory for four-and-a-half years (Strathmann and Strathmann, 2007). Pelagic dispersal duration of about a year is believed to be common among deep-sea invertebrate larvae that require extensive transport to the nearest site suitable for settlement (Adams et al., 2012). In 1973, Turner proposed that deep-sea larvae delay metamorphosis for long periods and remain in the benthic boundary layer until a suitable habitat is detected (Turner, 1973). This phenomenon may also apply to larvae that migrate to surface waters from methane seeps (Arellano et al., 2014) and from hydrothermal vents. Such plasticity may be essential for longterm species survival given the unpredictability of the oceanic environment.
The fact that during periods of nutritional hardship, echinoid larvae up-regulate both CREB and NFKB (Figs. 2 and 5) is interesting in light of recent studies showing that adult humans who experienced adverse socioeconomic circumstances as children display a highly sensitized, pro-inflammatory "defensive" phenotype marked by up-regulation of those same factors (Miller et al., 2009). This raises the possibility that genomic encodings that mediate developmental programming in response to early-life adversity may have arisen early in metazoan evolution and be widely conserved among animal species.
We thank Steve Eddy of the University of Maine CCAR for culturing and providing larvae and algae, Timothy Stearns of MDI Biological Laboratory for Gene Ontology enrichment analysis, and Andy Cameron of Caltech for helpful comments on the manuscript prior to submission. Research reported in this publication was supported by the NSF research experience for undergraduate (REU) program at the MDI Biological Laboratory (DBI 0453391), and by Institutional Development Awards (IDeA) from the National Institute of General Medical Sciences of the National Institutes of Health under grant numbers P20-GM104318 and P20-GM103423.
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TYLER J. CARRIER (1,2), BENJAMIN L. KING (1), AND JAMES A. COFFMAN (1*)
(1) MDl Biological Laboratory, Salisbury Cove, Maine 04672; and (2) School of Marine Sciences, University of Maine, Orono, Maine 04469
Received 14 November 2014; accepted 9 April 2015.
(*) To whom correspondence should be addressed. E-mail: email@example.com
Table 1 Sequences of primers used in qRT-PCR Gene Forward Primer Reverse Primer Sd-FoxO TCATTCTGGACGCGGACAAA ATCAGCGCTTGACCTTGTCA Sd-4EBP CAGGCCTCTGTGAGCATTCA ACGAGAAAGAGCTGCCGAAA Sd-ATPsyn CTTGCATTGGGTTCATGCCA ATTGGGGCGGAGTTTCTCTG Sd-CytC ATGCTGCCCAATGTGTTTTT GAATGCTTGTGTGTCGGAGA Sd-Nuc TGTCGGACATTTTGTTGAAGA TTTTTGTGTATGTCAGTTGCATAAT Sd-HSPE1 ATATGCGACCACAGCCAGAG CAGCCGTTTGCAAGACAGTG Sd-NFkB TGCCCAGGTTACAGCTAACG AGAGAAGCGCATGTGTCACA Sd-CREB AAGACAGCCAAGGGAATCCC AACTTCTGCTGCTCGACTCC Sd-EAAT AAACAGGAAAGCCTGGCATA GACTTGAGATGGGCAGCAAT Sd-FoxJ1 TGGCAGAATTCCATCCGTCA AGGCGTGTCGTCTCTTCTTG Sd-Elk TTCATTGGCCCGCCATTTTG CCGACCCGCCATTTCGTATA Sd-HPRT CTCAACTGGAGGTCAACCCC AAGTTGGCTTTCTGGACCCC
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|Author:||Carrier, Tyler J.; King, Benjamin L.; Coffman, James A.|
|Publication:||The Biological Bulletin|
|Date:||Jun 1, 2015|
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