Metabolic profiling of late-term turkey embryos by microarrays

MOLECULAR, CELLULAR, AND DEVELOPMENTAL BIOLOGY Metabolic profiling of late-term turkey embryos by microarrays J. E. de Oliveira,*1 S. Druyan,*2 Z. Uni,† C. M. Ashwell,* and P. R. Ferket*3 *North Carolina State University, Prestage Department of Poultry Science, Raleigh 27695; and †Hebrew University, Faculty of Agriculture, Department of Animal Science, Rehovot, Israel 76100 tase, acetyl-CoA carboxylase, lipoprotein lipase, and thyroxine deiodinase) had reduced expression between E22 and E26, corresponding to the period of expected limited oxygen supply. In contrast, genes related to opposing pathways in carbohydrate metabolism, such as glycolysis and gluconeogenesis (hexokinases, glucose-6 phosphatase, phosphofructokinases, glucose 1–6 phosphatase, pyruvate kinase, and phosphoenolpyruvate carboxykinase), or glycogenesis and glycogenolysis (glycogen synthase and glycogen phosphorylase) had rather static expression patterns between E22 and E26, indicating their enzymatic activity must be under posttranscriptional control. Metabolic survey by microarray methodology brings new insights into avian embryonic development and physiology.

Key words: poultry embryo, turkey, gene expression, microarray, metabolism 2013 Poultry Science 92:1011–1028 http://dx.doi.org/10.3382/ps.2012-02354

INTRODUCTION Poor hatchability and fitness of hatchling poults adversely affects production efficiency and animal welfare in the turkey industry. Hatchability has not significantly improved during the last 20 yr (Schaal and Cherian, 2007), and weak poults and “starveouts” are also common problems associated with commercial turkey production (Christensen et al., 2003). High embryonic mortality has been shown to occur between internal pipping and egg emergence (Christensen et al. 2003), but little is known about the metabolic constraints and patterns of change during the late stages of incubation. Since the completion of the chicken genome (International Chicken Genome Sequencing Consortium, 2004), new opportunities have become available for the study of avian biology (Lamont, 2006), particularly during ©2013 Poultry Science Association Inc. Received March 28, 2012. Accepted December 7, 2012. 1 Current address: Cargill R&D Centre Europe, Havenstraat 84, B1800 Vilvoorde, Belgium. 2 Current address: Institute of Animal Science, Agricultural Research Organization, The Vulcani Center, PO Box 6, Bet Dagan 50250, Israel. 3 Corresponding author: [email protected]

the critical period of perinatal development. Even though turkey sequences are just now becoming available for most genes, it has been demonstrated that gene sequences of chickens and turkeys are similar enough to use chicken probe sequences to hybridize with turkey target RNA (Reed et al., 2005; De Oliveira et al., 2009). Investigation of gene expression during critical periods of development could accelerate the identification of metabolic distress points that affect early growth performance and viability. Complex biological systems that undergo sudden shifts, as observed in the energy metabolism of lateterm avian embryos, are good candidates for highthroughput DNA microarray studies (Spielbauer and Stahl, 2005). Previous molecular approach methods were very time consuming, expensive, and subject to many causes of variation (Harris, 2000; Wiseman, 2002; Yadetie et al., 2004). Most genes involved in energy metabolism have well-known sequences, so a 2-color focused microarray could be designed with selected genes, where statistical power can be increased by replication and all genes could be annotated. This approach could greatly increase the ability to detect even small changes in gene expression by taking advantage of replication when building a general ANOVA

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ABSTRACT The last stages of embryonic development are crucial for turkeys as their metabolism shifts to accommodate posthatch survival and growth. To better understand the metabolic change that occurs during the perinatal period, focused microarray methodology was used to identify changes in the expression of key genes that control metabolism of turkey embryos from 20 d of incubation (E) until hatch (E28). Gene expression patterns were evaluated in liver, pectoral muscle, and hatching muscle and were associated with measured embryonic growth and tissue glycogen concentration. Within the studied period, the expression of 60 genes significantly changed in liver, 53 in pectoral muscle, and 51 in hatching muscle. Genes related to lipid metabolism (enoyl-CoA hydratase, 3-hydroxyacyl-CoA dehydrogenase, 3-hydroxymethylglutaryl-CoA reduc-

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MATERIALS AND METHODS Egg Incubation and Embryo Tissue Sampling Two-hundred fertilized turkey eggs were obtained from a commercial hatchery (Prestage Farms, Clinton, NC), divided into groups with similar weight distribution (74 ± 10 g), and incubated under standard conditions. Twenty-four eggs were sampled at embryonic age (E) 20, E22, E24, E26, and at day of hatch (E28). Eggs were opened to extract the embryos, which were immediately euthanized by cervical dislocation. Liver, pectoral muscle, and hatching muscle was removed from each embryo, and a section from each tissue was placed in RNA Later (Ambion Inc., Austin, TX) solution for subsequent RNA isolation. To associate gene expression data with poult energy status, the remaining liver, pectoral muscle, and hatching muscle tissues were immediately placed on ice and subsequently stored at −20°C for determination of tissue glycogen concentration [GLY]. Glycogen was extracted from approximately 1 g of tissue sample, and [GLY] was assayed with reagents within 30 min of preparation to minimize technical error of the modified iodine binding method (Foye et al., 2006) described by Bennett et al. (2007). The experimental protocol was approved by the North Carolina State University Institutional Animal Care and Use Committee.

Microarray Database Accession Numbers Minimal information about the microarray platform, experiment protocols, and microarray gene expression data has been deposited in the publicly available NCBI GEO Archive (http://www.ncbi.nlm.nih.gov/geo/) under microarray platform accession number GPL6041.

Time course data accession numbers are GSE9399 for liver, GSE9472 for breast muscle, and GSE9473 for the hatching muscle data.

RNA Isolation and Microarray Hybridization The RNA was extracted from 100 mg of tissue using TRI Reagent (Molecular Research Center Inc., Cincinnati, OH). Tubes were filled with 0.3 mL of 0.1-mm glass beads and shaken in a Mini-Beadbeater-96 (Biospec Inc., Bartlesville, OK) until the sample tissue was completely homogenized. Isolated RNA was quantified using a ND-1000 spectrophotometer (NanoDrop Technologie Inc., Wilmington, DE), and RNA integrity was verified by electrophoresis on 1.5% agarose gel. Equal amounts of RNA extracted from samples acquired from 6 replicate animals on the same embryonic day were pooled and adjusted to 0.5 µg/µL of concentration, resulting in 4 biological replicates that were indirectly labeled, 2 with Cy3 and 2 with Cy5, according to the experimental design (Figure 1). On each array, a combination of 33 pmol of cDNA labeled with different CyDyes were hybridized for 16 h at 42°C. Microarray probes consisting of 70-mer bp oligonucleotides from 90 selected Gallus gallus genes (TIGR, 2004) were printed on amino-silane coated slides (UltraGAPS Coated Slides, Corning Inc., Acton, MA). Each oligonucleotide sequence was printed twice in each grid, with grids repeated 4 times on a slide, totaling 8 replicate spots per slide. Printed genes included 41 genes from carbohydrate metabolism (glycolysis, gluconeogenesis, tricarboxylic acid cycle (TCA), and pentose phosphate pathways), 13 genes from glycogenesis and glycogenolysis pathways, 11 genes from lipid metabolism, 18 genes from hormone related metabolism, and 7 intestinal enzymes and nutrient transporter genes. Six more probes were added, including 3 housekeeping genes (GAPDH, chßact, and chEF2), one plant gene (atCAB2-arapdopsis), and 2 repeats of turkey genomic DNA to serve as internal controls. Thus, each array had a total of 96 probes on the array. A list of relevant genes present in the array was previously published (De Oliveira et al., 2009). Slides were scanned on a ScanArray GX PLUS Microarray Scanner (PerkinElmer Life and Analytical Sciences, Shelton, CT) set to 65% laser power.

Array Data Processing and Statistical Analysis The experimental design was a complete interwoven loop (Garosi et al., 2005) as presented in Figure 1. Spot quantification was carried out using ScanAlyze Software (Eisen et al., 1998). Raw data files were joined, transformed to a log2 base, and analyzed in JMP Genomics (SAS Institute, 2007). Array data were analyzed using the normalization method (Wolfinger et al., 2001) based on the overall ANOVA method (Kerr et al., 2000), which uses the logarithms of the original

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normalization model for the logarithms of original fluorescence measurements, as previously described (Kerr et al., 2000; Wolfinger et al., 2001; Druyan et al., 2008). Moreover, recent developments in system biology allow for the construction of maps that display the interaction among genes, proteins, and transcription factors into protein-protein, signaling, metabolic, and transcription-regulatory networks (Brazhnik et al., 2002; Hassan et al., 2007), so visualization of these complex data can facilitate understanding of developmental and physiological changes. A previous study demonstrated the power of this approach in the study of intestinal maturation in turkeys (De Oliveira et al., 2009). The objective of this study was to apply the same technique to identify subtle changes in gene expression in liver and muscle that influence physiology during the critical period from internal pipping to hatch, and associate this information with measured embryonic growth and energy status. This information would be useful to better understand perinatal development and identify consequences of in ovo or incubation manipulation.

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fluorescence measurements instead of the log ratio values. This previously described model (Druyan et al., 2008) accounts for experiment-wide systematic effects that could bias inferences made on data from individual genes, which is a better way to analyze data with complex experimental designs and a minimum of 3 replicates per spot (Wolfinger et al., 2001). The residuals from this model were analyzed by mixed ANOVA in JMP Genomics (SAS Institute, 2007) according to the gene-specific model: Y = µ + E + Dye + Hyb + Slide + e,

Embryonic Growth and [GLY] Statistical Analysis Embryonic growth and [GLY] followed the same completely randomized experimental design. Data were analyzed by fit model procedure in JMP (SAS Institute. 2005), according to the model Y = µ + E + e, where day of incubation (E) was the fixed effect. Treatment means were compared by Tukey’s test (P < 0.01).

Metabolic Maps To interpret the metabolic mechanisms and their change as the embryo develops, a list of relevant genes present in this microarray was used to create a customized pathway map using Pathway Map Creator from Thomson Reuters (Thomson Reuters, London, UK). The gene list was produced with an annotation file including human gene symbols for each cor-

Figure 1. Experimental design. Complete interwoven loop design with 4 biological replicates per time point. Each arrow indicates one microarray slide. Filled arrow bottom indicates sample labeled with Cy3, and open arrow tip indicates sample labeled with Cy5. E = day of incubation.

responding chicken gene. To obtain an overall view of expressed genes and pathway flux, all intensity standardized means were overlaid on the pathway map using MetaCore (Thomson Reuters) to generate figures (maps) containing gene expression representation for each studied embryonic age and tissue. Genes with statistically significant differences in expression (P < 0.01) are circled on each map (Figures 3, 5, and 7). A list of gene abbreviations and their corresponding names can be found in Appendix Table A1. A list of symbols used on the maps is provided in Appendix Figure A1.

RESULTS AND DISCUSSION Embryonic Development and Energy Status The weight of embryos and sampled tissues are presented in Table 1. Embryonic yolk-free BW (YFBW) increased at an average rate of 4.6 g/d from E20 until hatch (E28; Table 1). This fast rate of growth was expected during the last stage of the developing turkey embryo (Moran, 2007), and part of the weight gain during the last 48 h (E26 to E28) is expected to be water transferred to embryo tissues from other egg compartments (Amos, 1991). During the same period, the yolk sac content was used, disappearing at an average rate of 1.9 g/d as embryo weight increased (Table 1). Pectoral muscle and liver tissues weights also increased daily from E20 until hatch (E28), with the most significant increase during the last 48 h before hatch (E26 to E28; Table 1). Hatching muscle growth could be divided in 3 distinct phases: E20 to E24, E24 to E26, and E26 to E28 (Table 1). The first phase describes hatching muscle growth before preparation for pipping (E20 to E24), with growth rate averaging 0.09 g/d. Between E24 to E26, hatching muscle growth rate increased 3.5-fold in preparation for internal pipping. The hatching muscle stopped growing after E26 as it reached its maximum size by the time of pipping and hatch (Table 1). During

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where Y = hybridization intensity, μ = mean intensity, and e = random error, with day of incubation (E) and Cy-Dye (Dye) as fixed effects and hybridization batch (Hyb) and Slide as random effects. Mean intensities were compared using false discovery rate at P < 0.01. Results were used to produce clustering plots utilizing hierarchical clustering, with distances between clusters defined by the Ward method (SAS Institute. 2005), with time as a nominal parameter (distances = 0). To facilitate interpretation of gene expression measurements, the expression pattern clusters within each tissue were identified by a distinct color and uppercase letter. This microarray platform was designed to reduce false-positive results and to pick up small differences in gene expression by greatly increasing replication and statistical power (Druyan et al., 2008). The platform has been tested, and the results of validation by PCR showed it to be more precise than that technique (Druyan et al., 2008; De Oliveira et al., 2009). More details about this microarray platform approach and statistical analysis have been previously reported (Druyan et al., 2008).

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Table 1. Turkey embryo organ development in the period between embryonic (E) d 20 and hatch (E28) Item Embryonic age (d)  E20  E22  E24  E26  E28 SEM Period (d)   E20   E22   E24   E26

to to to to

E22 E24 E26 E28

YFBW1 (g)

YFBW2 (%)

Yolk (g)

Pectoral muscle3 (%)

22.38e 29.23d 37.89c 43.16b 59.53a 0.486

30.61d 38.94c 50.98b 59.46a — 0.467

18.78a 18.97a 17.73a 15.15b 7.58c 0.336

25.73a 25.23ab 23.82b 20.87c 13.27d 0.447 Daily growth

3.43 4.33 2.64 8.185

4.0 6.0 4.2 —

— −0.62 −1.29 −3.79

— −0.71 −1.48 −3.80

Yolk/egg (%)

4.492bc 4.721ab 3.980d 4.212cd 5.037a 0.089 0.115 −0.366 0.116 0.413

Liver3 (%)

Hatch muscle3 (%)

1.253c 1.398b 1.467bc 1.495b 1.898a 0.045

0.306d 0.404d 0.664c 1.299a 1.136b 0.032

0.073 0.035 0.014 0.201

0.049 0.130 0.318 0.034

followed by different lowercase letters in a column are statistically different (P < 0.0001); n = 24. = embryo yolk-free BW. 2Embryo YFBW expressed as percentage of egg weight. 3Expressed as percentage of embryo YFBW. 1YFBW

this E20 to E26 period, the embryo also accumulates nutrients into tissues from other egg components, and general organ maturation is promoted (Moran, 2007). The hatching muscle development observed in this study follows a pattern previously described for chicken embryos (Romanoff, 1960; John et al., 1987; Moran, 2007). The results of glycogen analysis of liver, pectoral muscle, and hatching muscle are presented in Table 2. Glycogen is synthesized from glucose produced by gluconeogenesis using amniotic fluid amino acids as substrate (Muramatsu et al., 1990), and then it is used to fuel the hatching process (Donaldson and Christensen, 1991). The accumulation of glycogen and its use in these tissues by the avian embryo can be compared with an athlete that prepares to run a marathon (Foye, 2005), where high amounts of glycogen are stored before extreme anaerobic exercise. Therefore, enzymes associated with glycogen metabolism in liver and muscle tissues are expressed in parallel. A more in-depth discussion about the importance of glycogen within each tissue during each observation period is presented later. The results of the gene expression cluster analysis and the corresponding pathway maps can be seen, respectively, in Figures 2 and 3 for liver, Figures 4 and 5 for pectoral muscle, and Figures 6 and 7 for hatching muscle. The greatest number of genes that showed differences in expression by embryonic age was the liver (56 genes), followed by pectoral muscle (52 genes) and hatching muscle (49 genes). It is important to note that housekeeping genes, GAPDH (Figures 2, 4, and 6) and chEF2 (Figures 2 and 6), can change significantly with advancing embryonic age in association with the high rate of development. Proteomic analysis in chickens was observed to have high expression levels of GAPDH, KCRM, enolase, and LDHA during both embryonic and posthatch development (Doherty et al., 2004; Agudo et al., 2005). Even very small changes in color intensity can be significant for genes with very consistent sig-

nals, due to the high sensitivity of this array (Druyan et al., 2008). For example, a gene with constant zero color intensity (or background intensity) can be significant if an intensity higher than zero spikes at one time point. The reasons for this microarray platform sensitivity have been previously discussed (Druyan et al., 2008). This may be why atCAB2-Arabidopsis (Figures 4 and 6), which is not present in animal tissues and turkey genomic DNA (Figures 2 and 6), showed as significant in some clusters. This inherent error can occur when using a false discovery rate of 0.01, because one false positive or false negative gene every 100 genes is expected. For this reason, conclusions from single point shifts on gene expression should be avoided when using this microarray platform without first confirming it by an independent technique such as real-time PCR. Therefore, the following results and discussion will be based on gene expression patterns rather than single Table 2. Turkey embryo glycogen concentration in liver, pectoral muscle, and hatching muscle from E20 to E28 (hatch) Glycogen tissue concentration (mg/g) Pectoral muscle

Hatch muscle

Item

Liver

Embryonic age (d)  E20  E22  E24  E26  E28 SEM Period (d)

0.929c 0.490b 7.513a 0.585a 7.418a 0.594a 4.762b 0.436b 0.884c 0.071c 0.348 0.025 Glycogen balance (mg/g per

  E20   E22   E24   E26

to to to to

E22 E24 E26 E28

3.292 −0.048 −1.328 −1.939

0.048 0.005 −0.079 −0.183

— — 0.436a 0.520a 0.059b 0.035 d)

    0.042 −0.231

a–cMeans followed by different lowercase letters in a column are statistically different (P < 0.0001); n = 24.

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a–eMeans

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point gene expression changes. A general discussion of each cluster by tissue follows.

Liver

at lower levels between E22 to E26 (Figure 2 cluster d, and Figure 3) than at E20 and E28. The same response in gene expression was observed for thyroxine deiodinase (DIO1), which coverts T4 to T3, the active form of one important hormone involved in stimulating fat mobilization (Figure 2 cluster d). These results indicate that the liver mobilization of fat is reduced during the plateau stage of oxygen consumption until external pipping occurs. Carnitine O-palmitoyl transferase II (CPTII), the enzyme responsible for long-chain FA transport into the mitochondria, had increased gene expression during the same period (E22 to E26) as indicated in Figure 2 (cluster a) and Figure 3. Rather than interpreting this response as promoted β-oxidation, it is more likely in response to a state of fasting, as previously observed with chicken CPTI gene expression (Skiba-Cassy et al., 2007). Also, it has been previously determined that allosteric control of CPT enzymes by malonyl CoA is even more important than their gene expression (Eaton, 2002; Bartlett and Eaton, 2004; Skiba-Cassy et al., 2007). This is further confirmed by the expression pattern of another enzyme involved in the process, carnitine acetylase (CRAT), which is upregulated only after E26 (Figure 2 cluster e, and Figure 3). Based on the hepatic gene expression profile observed in this study, fat metabolism in the turkey embryo is inhibited between E22 to E26, which is when oxygen supply is expected to be most limited, but it does not prevent the poultry embryonic liver from continuing to accumulate lipids transferred from the yolk-sac (Pulikanti et al., 2010). Liver Glycogen Metabolism. Liver glycogen metabolism is very important to sustain avian embryos during hatch (Donaldson and Christensen, 1991; Donaldson et al., 1991; Moran, 2007). As discussed above, lipid metabolism is downregulated during the plateau stage of oxygen consumption and pipping, so the embryo must depend on anaerobic metabolism of carbohydrates for energy. Amnion consumption provides a substrate for the synthesis of glycogen that must be stored before hatching (Moran, 2007; De Oliveira et al., 2008). Liver glycogen concentration was observed to increase by 8-fold between E20 and E22 (Table 2), which is the same period in which the rate of amnion consumption by the embryo is greatest (De Oliveira et al., 2008). The liver is the main glycogen-storing organ, being able to store 5% of its weight as glycogen compared with 1% in muscle (Krebs, 1972). Our observations (Table 2) agree with those of other reports that liver glycogen concentrations in poultry embryos and hatchlings range between 1 and 3% of liver weight (Donaldson et al., 1991; Uni et al., 2005; Foye et al., 2006). Glycogen reserves mobilized during hatching yields glucose that can be exported to other tissues, which is reduced to lactate when oxygen is limiting (Lindy and Rajasalmi, 1966; De Oliveira et al., 2008). The rate of glycogen utilization was greatest in the liver, which was 6-fold greater than use rate in hatching muscle, and 15-fold greater than glycogen use rate

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The large number of significant genes (56 total) was expected for liver because it is the most metabolically active organ (Krebs, 1972), especially during this stage of development when the embryo is preparing for its posthatch life. The liver is known for being the organ with the most enzymes and able to perform almost all the metabolic pathways active in the animal (Kietzmann and Jungermann, 1997). During the last days of incubation, 2 main hepatic functions can be highlighted: 1) storage of energy as glycogen for hatching (Uni and Ferket, 2004), and 2) transfer of many nutrients from other egg compartments to liver, including large amounts of cholesterol from the yolk sac (Moran, 2007). The total amount of lipids in the chicken liver has been shown to increase from 24 mg at d 15 to 150 mg at 19 d of incubation (Noble, 1986; Pulikanti et al., 2010). The metabolic map of the liver is illustrated in Figure 3, where genes with statistically significant differences in expression are circled. Liver Lipid Metabolism. One of the main objectives of this study was to identify the stage of embryonic development when the embryo switches from lipid-based to carbohydrate-based metabolism. This metabolic shift has been suggested by several authors (Romanoff, 1967; Donaldson et al., 1991; Noy and Sklan, 1999; Uni and Ferket, 2004; Foye, 2005; Moran, 2007) because fatty acid (FA) β-oxidation is limited by low oxygen supply around the time of internal pipping stages, and the anaerobic catabolism of the stored glycogen ensues. Based on the gene expression patterns observed in this study, FA β-oxidation inhibition apparently starts around E22 (Figure 2) because enoyl-CoA hydratase (ECHDC1) and 3-hydroxyacyl-CoA dehydrogenase (HCD2), which are respectively responsible for the second and third steps of β-oxidation, were significantly downregulated after E22 (Figure 2 clusters b and d, and Figure 3). Other indications of reduced lipid metabolism after E22 are supported by the downregulation of 3-hydroxy-methylglutaryl-CoA reductase (HMDH), which is important for cholesterol synthesis, and acetylCoA carboxylase (ACACA), which is the first step of FA synthesis (Figure 2 cluster b, and Figure 3). These results are in agreement with earlier findings that suggest that even though the liver is packing and accumulating lipids during this stage, lipid processing occurs in the yolk sac membrane before it is transported to the liver (Noble, 1986). The acetyl-CoA acetyltransferase gene (ACAA1), for the enzyme responsible for removing acetyl-CoA units from FA during degradation, showed declining expression toward hatch (Figure 2 cluster c, and Figure 3). The gene for lipoprotein lipase (LPL), responsible for triacylglycerol hydrolysis into FA + glycerol during fat mobilization, was expressed

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Figure 2. Clustering of genes significantly affected by time in embryonic liver. Gene symbols appear on the left, with the color heat map indicating fold difference in the middle, and the clustering by pattern similarity on the right. Expression pattern groups are identified by lowercase letters (a, b, c, and d). E = day of incubation.

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Figure 3. Customized pathway map representing a summary of gene expression patterns observed in turkey embryonic liver, created using Pathway Map Creator (Thomson Reuters, London, UK). Map objects represent compounds, enzymes, reactions, and cell compartmentalization. Gene expression data are added as thermometers by the corresponding gene product. Thermometer numbers correspond to embryonic age, 1 = E20, 2 = E22, 3 = E24, 4 = E26, and 5 = E28, respectively. A complete legend to the symbols representing each gene is provided in Appendix Figure A1. Color version available in the online PDF.

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Figure 4. Clustering of genes significantly affected by time in embryonic pectoral muscle. Gene symbols appear on the left, with the color heat map indicating fold difference in the middle, and the clustering by pattern similarity on the right. Expression pattern groups are identified by lowercase letters (a, b, c, and d). E = day of incubation.

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Figure 5. Customized pathway map representing a summary of gene expression patterns observed in turkey embryonic pectoral muscle, created using Pathway Map Creator (Thomson Reuters, London, UK). Map objects represent compounds, enzymes, reactions, and cell compartmentalization. Gene expression data are added as thermometers by the corresponding gene product. Thermometer numbers correspond to embryonic age, 1 = E20, 2 = E22, 3 = E24, 4 = E26, and 5 = E28, respectively. A complete legend to the symbols representing each gene is provided in Appendix Figure A1. Color version available in the online PDF.

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Figure 6. Clustering of genes significantly affected by time in embryonic hatching muscle. Gene symbols appear on the left, with the color heat map indicating fold difference in the middle, and the clustering by pattern similarity on the right. Expression pattern groups are identified by lowercase letters (a, b, c, and d). E = day of incubation.

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Figure 7. Customized pathway map representing a summary of gene expression patterns observed in turkey embryonic hatching muscle, created using Pathway Map Creator (Thomson Reuters, London, UK). Map objects represent compounds, enzymes, reactions, and cell compartmentalization. Gene expression data are added as thermometers by the corresponding gene product. Thermometer numbers correspond to embryonic age, 1 = E20, 2 = E22, 3 = E24, 4 = E26, and 5 = E28, respectively. A complete legend to the symbols representing each gene is provided in Appendix Figure A1. Color version available in the online PDF.

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genesis (G6PT, F16P, and PPCKC) in the liver, the complexity is evident. These antagonistic pathways apparently work together to favor the flux of substrate toward other connecting pathways, such as glycogenesis, glycogenolysis, and the pentose-phosphate pathway. For example, the enzyme at the first point of control of liver glycolysis, glucokinase (HXK4), is upregulated on E28 (hatch; Figure 2 cluster e, and Figure 3), although the opposing gluconeogenic enzyme, glucose-6-phophatase (G6PT), is also upregulated at the same time (Figure 2 cluster e, and Figure 3). Other examples of apparent contradictory regulation include phosphoenolpyruvate carboxykinase (PPCKC), the point of control for gluconeogenesis, and pyruvate kinase (PKM2), the point of control for glycolysis. The PPCKC and PKM2 both have the same expression pattern (Figure 2 cluster e, and Figure 3). Instead of exhibiting conflicting pathway controls, this observation illustrates the liver’s ability to move substrates up and down these pathways during this critical period of embryonic development, and it demonstrates the importance of posttranscriptional control to regulate actual protein levels. High levels of some enzymes, such as HXK1 and TYR, close to hatching have been confirmed by proteomics in chicken (Jianzhen et al., 2007). Based on measured glycogen stores and utilization (Table 2), gluconeogenesis must also predominate until E26, and glycolysis must predominate afterward. Thus, expressed enzyme activities must be influenced by other forms of control, such as hormones, substrate/ product concentrations, and posttranslational modification. There were 2 isoforms of PFK present on the array; one is the muscle isoform (PFKM; Figure 2 cluster e, and Figure 3) and the other is the liver isoform (PFKL) (Figure 2 cluster a, and Figure 3). The PFKL appears in the same cluster as the insulin receptor gene (INSR; Figure 2 cluster a), which indicates by their expression patterns that these 2 gene products may work in concert to favor glycolysis between E22 and E26. The last enzyme considered as a significant point of regulation of liver gluconeogenesis is fructose 1–6 biphosphatase (F16P or F16Q), which was downregulated after E22 until hatch (Figure 2 cluster b, and Figure 3). This observation is further evidence that gluconeogenesis predominate before pipping, whereas glycolysis is favored to allow utilization of glucose produced by glycogenolysis only between E22 and E26, as demonstrated by peaks of HXK4 and INSR. In general, the hepatic gene expression results observed in this study confirmed current knowledge of the liver’s role in supporting energy of other tissues by switching energy metabolism from different substrates, depending on oxygen supply (i.e., from lipids to carbohydrates and protein), to fuel increased physical activity during hatching. The major contribution of these findings is to pinpoint exactly when these events occur during late-term embryonic development.

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in pectoral muscle (Table 2). The rate of glycogen use in hatching muscle was 2.5-fold greater than in pectoral muscle (Table 2). These differences in glycogen use rate among tissues confirm the hypothesis that the liver must supply glucose to maintain energy homeostasis of the whole organism (Krebs, 1972), and validates the use of glycogen to fuel the intense activity of hatching muscle during pipping and hatching in comparison with the participation of the pectoral muscles in emergence from the shell. By associating data from tissue growth with glycogen concentration, an intricate pattern of physiological events can be seen. It is clearly evident why turkey embryos are so sensitive to incubation distress during this critical period of embryonic development. If tissue growth, energy stores, or energy use is compromised, the embryo may not be able to emerge from the shell and may die because of metabolic energy constraints. The 2 main enzymes of glycogen metabolism are glycogen synthase (GYS1) and glycogen phosphorylase isoforms (PYGL and PYGM), which are, respectively, responsible for synthesis and mobilization of glycogen (Onoagbe, 1993; Carrizo et al., 1997; Seki et al., 2005). The glycogen synthase gene (GYS1) was upregulated between E26 and E28 (Figure 2 cluster e), whereas glucose-6-phosphate isomerase (G6PI), which connects glycolysis to glycogen pathways, was upregulated between E22 and E26 (Figure 2 cluster a). Increasing hepatic GYS1 activity has been reported to occur during chicken late-term embryonic development until hatch (Carrizo et al., 1997), which is in agreement with gene expression data presented herein. This is the period when internal and external pipping is occurring. As expected, glycogen synthase kinase-3-β (GSK3B), which inhibits GYS1 (Grimes and Jope, 2001), is downregulated after E22 until hatch (Figure 2 cluster b). The liver isoform of the glycogen phosphorylase gene (PYGL) was upregulated from E24 until hatch (Figure 2 cluster e, and Figure 3) in parallel with GYS1. This observation confirms that glycogen phosphorylase is needed to mobilize glycogen reserves starting at the time of internal pipping (Table 2). Based on enzyme gene expression found in this study, glycogen synthesis predominantly occurs between E24 and E26, whereas glycogen degradation predominantly occurs after E26. These observations are supported by the measured liver glycogen concentration (Table 2). Creatine phosphate is another possible energy source that can be produced in chicken liver (Zhan et al., 2006). Creatine phosphate is produced by the enzyme creatine kinase (KCRM), which was downregulated during the period when glycogen metabolism is most active (E22 to E26) as shown in Figure 2 (cluster d) and Figure 3. Thus, this energy source may not be very important in the liver during this period of embryonic life. Liver Glycolysis/Gluconeogenesis. Observing the expression of key enzymes that control flux favoring glycolysis (HXK1, PFKL, and PFKM) or gluconeo-

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Pectoral Muscle

(Figure 4 cluster b, Figure 5), probably because of glucose coming into the pathway through glycogenolysis (Table 2). Most TCA cycle enzymes, such as citrate synthase (CS), isocitrate dehydrogenase (IDH2 and IDH3A), and malate dehydrogenase (MDH1), showed similar downregulated gene expression patterns (Figure 4, cluster d) from E20 until E26, which was expected because limited oxygen supplies cause glycolysis to produce lactate instead of acetyl-CoA. Exceptions to this observation were the 2 succinate enzymes, succinate-CoA ligase (SUCLG1) and succinate dehydrogenase (SDHB), which were upregulated until E26, with a further upregulation between E26 and hatch (Figure 4 cluster b, Figure 5). This is an interesting finding because catabolism of branch-chain amino acids is common in muscle, and their carbon backbone enters the TCA through succinate (Murray et al., 2003). Increased succinate metabolism may be an indication of pectoral muscle protein degradation to provide energy, which is especially important at hatch as suggested by several authors (Donaldson and Christensen, 1991; Christensen et al., 2003; Uni and Ferket, 2004; Foye, 2005; Moran, 2007).

Hatching Muscle The specific function of the hatching muscle is to empower the neck to break the egg membranes and the shell during pipping and hatching, so the most critical period of activity for this muscle is through hatch (Moran, 2007; De Oliveira et al., 2008) as the poult’s ultimate survival depends on it. The 4 clusters of gene expression patterns in the hatching muscle are illustrated in Figure 6, and its metabolic map is presented in Figure 7. Hatching Muscle Growth. Some growth-related genes in hatching muscle changed significantly during the prenatal period. These genes were the growth hormone-inducible soluble protein (LYRM5; Figure 6, cluster b), which showed peak upregulation at E22, and the thyroid hormone receptor α (TR-α; Figure 6 cluster d, Figure 7), which was downregulated after E26. The growth pattern of the hatching muscle presented in Table 1 shows rapid growth right before full mass of the hatching muscle was achieved at E26. This rapid increase in muscle mass is a consequence of cell hyperplasia and hypertrophy, but also due to lymph infiltration (Fisher, 1958; Pulikanti et al., 2010; Sokale et al., 2011). These muscle development events coincide with expression of the TR-α gene (Figure 6 cluster d, Figure 7), indicating that thyroid hormones may play an important role in the growth and maturation of the pipping muscle. Thyroid hormones have been previously associated with perinatal maturation of turkeys (Christensen et al., 1996, 2003). Similar observations were made in chickens, where the hatching muscle increases in size until it accounts for more than 1% of the embryo’s body (Sokale et al., 2011), and it also accu-

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The importance of the pectoral muscle during lateterm embryonic development in birds is related its relatively large size, and its possible function as glucogenic energy store to be used upon demand (De Oliveira et al., 2008). The 4 clusters of gene expression patterns in pectoral muscle can be found in Figure 4, and its metabolic pathway is presented in Figure 5. Pectoral Muscle Growth. Three genes associated with growth characteristics were present in Figure 6 cluster a: growth hormone inducible-soluble protein (LYRM5), growth hormone and its receptor (GH and GHR), indicating a peak in expression 48 h before hatch (E26; Figure 4 cluster a, Figure 5). Thus, measured pectoral muscle mass increased continually throughout the study period (Table 1). The expression of genes involved in growth peaked at E26, followed by their subsequent downregulation (Figure 4 cluster a). This observation agrees with a previous report that turkey embryos reach full body size around E26, with posterior increase in muscle mass toward hatch being a consequence of increased tissue hydration (Vleck, 1991). Pectoral Muscle Glycogen Metabolism. As observed in liver, genes for isoforms of glycogen synthesis (GYS1) and glycogen mobilization (PYGM) in pectoral muscle were also present in the same cluster (Figure 4, cluster b), indicating that both enzymes are present and their functionality probably depends on posttranslational forms of control, such as phosphorylation (Grimes and Jope, 2001). As shown in Table 2, pectoral muscle glycogen concentration decreased from E24 until hatch. Because muscle tissue cannot export glucose, a decrease in [GLY] indicates that pectoral muscle is utilizing glucose to support its activity during pipping and hatching. Pectoral Muscle Energy Utilization. Expression of genes associated with lipid metabolism in the pectoral muscle revealed that 3 enzymes involved in FA β-oxidation, carnitine O-palmitoyl transferase (CPTII), enoyl-CoA hydratase (ECHDC1), and thyroid hormone receptor α (TR-α), were downregulated after E24 until hatch (Figure 4 cluster c, Figure 5). However, 3 other enzymes of the same pathway, acetyl-CoA acyltransferase (ACAA), 3-hydroxyacyl-CoA dehydrogenase (HCD2), and carnitine acetylase (CRAT) were upregulated after E26 (Figure 4 cluster b, Figure 5). Combining these patterns of change in gene expression led to our conclusion that FA oxidation in pectoral muscle decreases after E22, as observed in the liver, but it becomes active again after external pipping when oxygen supply increases. The glycolytic enzymes, hexokinase (HXK1) and phosphofructokinase-1 (PFKM), are in clusters with opposite patterns from E20 until E26 (Figure 4, cluster b vs. cluster d). Even though pectoral muscle uptake of glucose was being reduced by lower HXK1 gene expression (Figure 4 cluster d, Figure 5), glycolysis was active as demonstrated by increased PFKM gene expression

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mulates proteins, glucose, and glycogen at the expense of fat in preparation for hatch (Pulikanti et al., 2010). Hatching Muscle Glycogen Metabolism. As in liver and pectoral muscle, enzymes of glycogen synthesis (GYS1) and degradation (PYGM) appeared in the same gene expression cluster (Figure 6 cluster a). Glycogen-related genes had higher expression between E24 and E26. The higher expression of glycogen synthase kinase-3 β (GSK3B) starting on E24 and toward hatch (Figure 6, cluster b), associated with the reduced expression of glucose-6-phosphate isomerase (G6PI) at E26 (Figure 6 cluster a, Figure 7), indicate the embryo’s need to inhibit glycogen synthase by phosphorylation, and the need to transport intermediates via the glycolysis pathway. Changes in hatching muscle glycogen concentration over time (Table 2) indicate that glycogen begins to be mobilized after E26, which may be a consequence of GYS1 inhibition (Figure 6, cluster a) by increased expression of GSK3B (Figure 6 cluster b). From the study of gene expression of enzymes related to glycogen synthesis and degradation in liver and muscle, we are apparently able to monitor but not predict pathway flux because both enzymes systems seem to be highly expressed during late-term incubation. So, the pathway flux associated with glycogen metabolism is likely regulated at the protein level by posttranslational modifications. Hatching Muscle Energy Utilization. Even though the contribution of the hatching muscle to break the shell is given mostly by hydraulic pressure of the turgid muscle rather than by muscle contraction (Smail, 1964), it has been demonstrated that energy is stored and used extensively by this tissue during the hatching process (De Oliveira, 2007; Pulikanti et al., 2010). Hatching muscle energy utilization shows a unique array of enzyme gene expressions, with FA β-oxidation enzyme 3-hydroxyacyl-CoA dehydrogenase (HCD2), glycolysis pathway enzyme pyruvate kinase (PKM2), and the succinate metabolism enzymes, succinate-CoA

ligase (SUCLG1) and succinate dehydrogenase (SDHB), all in the same cluster (Figure 6 cluster a). Their gene expressions were all upregulated between E24 and E26 when energy demand of this tissue would be at its maximum due to pipping activity. This observation leads us to conclude that the hatching muscle may be able to simultaneously use glycogen, FA, and amino acids as energy sources, which was not observed in any other tissue in this study. Even the TCA cycle enzymes, CS and isocitrate dehydrogenase (IDH3A), were upregulated in hatching muscle at E24 (Figure 6 cluster b) when oxygen supply is most limiting and hatching muscle is working during internal pipping. The spike in gene expression of TCA-related enzymes (CS, ME1, IDH3A, and GLUD1) at hatch was expected because oxygen supply was restored upon external pipping. Because distinct types of muscle fibers are found in the hatching muscle (John et al., 1987), it is possible that enzymes of opposing or different pathways are also being expressed in different muscle fiber types. Proteomic analysis of chicken hatching muscle before lymph infiltration found proteins related to 690 different cellular functions, a great part involved in metabolic processes, regulation of biological processes, and transport proteins (Sokale et al., 2011), which confirms the complexity here observed in the tissue transcryptomics. The results of this study can be summarized into distinct phases of events as illustrated in Figure 8. Embryonic growth rate is greatest at E20, fueled by yolk lipids. The embryo reaches its full size at E22 when oxygen demand is highest, but then oxygen supply starts to become limiting, so metabolism switches from using lipid to carbohydrate as the preferred energy source. Glycogen reserves are maximized at E22, but subsequently the balance between glycogen synthesis and degradation becomes negative, causing a transient reduction in tissue glycogen concentration. Hatching muscle is unique among other tissues in that it has all the energetic pathways operating at once. From E24

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Figure 8. Summary of events happening during late-term turkey embryo development.

TURKEY EMBRYO METABOLIC PROFILING BY MICROARRAY

until hatch, hormones peak and pipping of the shell progresses until the hatching process is completed.

Conclusion

ACKNOWLEDGMENTS This research was founded by Embrex Inc. (Durham, NC) through the USDA Small Business Innovation Research (SBIR) Phase II grant 2005-33610-16491, September 15, 2005, to September 14, 2007.

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Through association between embryonic growth measurements and gene expression patterns obtained by microarrays, it was possible to identify and confirm the pattern of expression of many key genes of turkey embryo metabolism and development. Our main finding was that the turkey embryo switches from lipid- to carbohydrate-based metabolism at 22 d of incubation, and that the enzymes related to glycogen metabolism in liver and muscle tissues are expressed in parallel. The approach used in this study could bring substantial contribution to the current knowledge about late-term embryo development and physiology in birds, particularly turkeys.

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APPENDIX Table A1. List of genes with respective gene symbols (abbreviations) Gene symbol

Gene name

ACAA1 ACACA ACADS ACLY ADH5 AK1 ALDH2 ALDOB ALDOC AMY2A ANPEP atCAB2 chEF2 CPTII CRAT CS DIO1 ECHDC1 ENO1 FH F16P G3P2 G6PI/T GAB3 GALT GAPDH GBE1 GHR GH GHRH

Acetyl-CoA acyltransferase Acetyl-CoA carboxylase Butyryl-CoA dehydrogenase Citrate lyase Formaldehyde dehydrogenase Adenylate kinase Aldehyde dehydrogenase Fructose biphosphate aldolase B Fructose-biphosphate aldolase α-Amylase Aminopeptidase Ey atCAB2-Arapdopsis chEF2-elongation factor 2 Carnitine O-palmitoyltransferase Carnitine acetylase ATP citrate synthase Thyroxine deiodinase I Enoyl-CoA hydratse/3-hydroxyacyl-CoA dehydrogenase Phosphopyruvate hydratase Fumarate hydratase Fructose 1,6 biphosphatase (F1,6 BPase) Glycerol-3-phosphate dehydrogenase Glucose-6-phosphate isomarase Aconitate hydratase Galactose-1-phosphate uridyltransferase Glyceraldehyde 3-phosphate dehydrogenase Glycogen branching enzyme Growth hormone receptor Growth hormone Growth hormone-releasing hormone Continued

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whole-body protein synthesis of the chicken embryo during incubation. Br. Poult. Sci. 31:101–106. Murray, R. K., D. K. Granner, P. A. Mayes, and V. W. Rodwell. 2003. Harper’s Biochemistry. 26th ed. McGraw-Hill Companies, Columbus, OH. Noble, R. C. 1986. Lipid metabolism in the chick embryo. Proc. Nutr. Soc. 45:17–25. Noy, Y., and D. Sklan. 1999. Different types of early feeding and performance in chicks and poults. J. Appl. Poult. Res. 8:16–24. Onoagbe, I. O. 1993. Hormonal control of glycogenolysis in isolated chick embryo hepatocytes. Exp. Cell Res. 209:1–5. Pulikanti, R., E. D. Peebles, R. W. Keirs, L. W. Bennett, M. M. Keralapurath, and P. D. Gerard. 2010. Pipping muscle and liver metabolic profile changes and relationships in broiler embryos on days 15 and 19 of incubation. Poult. Sci. 89:860–865. Reed, K. M., T. P. Knutson, S. B. Krueth, L. R. Sullivan, and L. D. Chaves. 2005. In silco mapping of ESTs from the turkey (Meleagris gallopavo). Anim. Biotechnol. 16:81–102. Romanoff, A. L. 1960. The Avian Embryo; Structural and Functional Development. Macmillian, New York, NY. Romanoff, A. L. 1967. Biochemistry of the avian embryo; a quantitative analysis of prenatal development. Interscience Publishers, New York, NY. SAS Institute. 2005. JMP Version 6 User’s Guide. SAS Inst. Inc., Cary, NC. SAS Institute. 2007. JMP Genomics Version 3.0 User’s Guide. SAS Inst. Inc., Cary, NC. Schaal, T., and G. Cherian. 2007. A survey of the hatchability of broiler and turkey eggs in the United States from 1985 through 2005. Poult. Sci. 86:598–600. Seki, Y., K. Sato, and Y. Akiba. 2005. Changes in muscle mRNAs for hexokinase, phosphofructokinase-1 and glycogen synthase in acute and persistent hypoglycemia induced by tolbutamide in chickens. Comp. Biochem. Physiol. B Biochem. Mol. Biol. 142:201–208.

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Table A1 (Continued). List of genes with respective gene symbols (abbreviations) Gene name

GHRHR GLUD1 GLUL GSK3B GYS1 HAGH HCD2 HMDH HXK1/4 IDH2 IDH3A INSR KCRM LDHA LDHD LPL LYRM5 MDH1 MDHM ME1 ME2 MGAM OGDH P2X1 PYC PEPT1 PFKL PFKM PGD PHKB PKM2 PPCKC PYGL PYGM SDHB SI SLC5A1 SUCLG1 TALDO1 TKT TR-α Turkey DNA TYR UGDH XPNPEP1

Growth hormone-releasing hormone precursor Glutamate dehydrogenase Glutamine synthetase Glycogen synthase kinase-3 β Glycogen synthase 1 (muscle) Hydroxyacylglutathione hydrolase 3-Hydroxyacyl-CoA dehydrogenase 3-Hydroxy-3-methylglutaryl-CoA reductase Hexokinase 1/4 Isocitrate dehydrogenase (NADP+) Isocitrate dehydrogenase (NAD+) Insulin receptor Creatine kinase l-Lactate dehydrogenase d-Lactate dehydrogenase Lipoprotein lipase Growth hormone-inducible soluble protein Malate dehydrogenase Malate dehydrogenase—muscle Malic enzyme Malate dehydrogenase Maltase-glucoamylase Oxoglutarate dehydrogenase ATP-gated ion channel receptor Pyruvate carboxylase Peptide transporter Phosphofructokinase (PFK-1) 6-Phosphofructokinase (PFK-1) Phosphogluconate dehydrogenase Phosphorylase kinase B α regulatory chain Pyruvate kinase Phosphoenol pyruvate carboxylase (PEPCK) Glycogen phosphorylase Glycogen phosphorylase (muscle) Succinate dehydrogenase Sucrase-isomaltase intestinal Sodium/glucose cotransporter 1 Succinate CoA ligase Transaldolase Transketolase Thyroid hormone receptor α Turkey genomic DNA Tyrosinase UDP-glucose 6-dehydrogenase Aminopeptidase P1

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Gene symbol

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Figure A1. Reference list with detailed description of symbols used in pathway maps in Figures 3, 5, and 7. MetaCore (Thomson Reuters, London, UK). Used with permission. Color version available in the online PDF.

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