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PHYSIOLOGY, ENDOCRINOLOGY, AND REPRODUCTION |
Department of Animal Sciences, Purdue University, West Lafayette, IN 47907
2 Corresponding author: phester{at}purdue.edu
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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-linolenic (C18:3n-3) acids for growth and development over myristic (C14:0) and oleic (C18:1n-9) acids.
Key Words: L-carnitine • hatch rate • egg production • yolk lipid • yolk sac
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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Egg formation within the oviduct begins after release of an ovum from the largest F1 ovarian follicle. The released ovum is transported via the oviduct, where egg-white proteins, produced in the magnum, are absorbed by the ovum as it traverses this region for approximately 3 h. As the ovum continues its movement through the oviduct, most of its development time will occur in the shell gland, where it imbibes nearly 15 g of water, CaCO3, and other components essential for shell formation (Etches, 1996). Because L-carnitine (β-hydroxy-
-trimethylammonium butyrate) plays a vital role in transporting long-chain fatty acids across the inner mitochondria membrane to produce energy through β-oxidation (Bremer, 1983), and because dietary L-carnitine can be intestinally absorbed in chickens (Duran et al., 2002), supplementation of L-carnitine may accelerate yolk lipid deposition (Peebles et al., 2007) and thus expedite follicular development. It may also increase metabolic rate in the magnum and shell gland and therefore affect albumen deposition and shell calcification, leading to greater egg weight and shell thickness.
All nutrients needed for embryogenesis are provided by the hen by the time the fertile egg is laid. In the United States, hen diets are composed mainly of corn and soy, which contain low levels of L-carnitine (Buyse et al., 2001). Therefore, eggs contain little or no L-carnitine (Chiodi et al., 1994). L-Carnitine is naturally synthesized in most animal species, including birds. Although L-carnitine biosynthesis increases during embryonic development, its levels are still much lower than those measured in adults because of the low activity of -butyrobetaine hydroxylase, the essential enzyme that catalyzes -butyrobetaine to L-carnitine (Borum, 1983; Rebouche, 1992). Yolk lipids provide essential energy to growing embryos. In fact, approximately 90% of the total energy requirement of the developing embryo is derived from fatty acid oxidation of yolk lipids (Noble and Cocchi, 1990). L-Carnitine also possesses antioxidant properties. Chick embryonic tissues contain high levels of polyunsaturated fatty acids, an essential component of the phospholipid content in cell membranes (Noble and Cocchi, 1990; Speake et al., 1998). Polyunsaturated fatty acids are susceptible to lipid peroxidation caused by free radicals, which are produced by mitochondria because of the high metabolic rate of rapidly developing embryos (Surai, 1999). L-Carnitine may work as an antioxidant to scavenge free radicals (Vicari and Calogero, 2001; Agarwal and Said, 2004; Agarwal et al., 2005). Thus, the presence of L-carnitine in the fertile egg may decrease embryonic mortality by reducing oxidative stress during the hatch process, thereby increasing hatch rate.
Previous work in our laboratory has shown that dietary supplementation of 125 ppm of L-carnitine, as compared with control diets, increased sperm concentration from 4.8 to 5.3 x 109 sperm/mL in White Leghorns (Zhai et al., 2007). Supportive data suggest that the mode of action of L-carnitine in increasing sperm concentration is through its antioxidant properties, thus extending the life span of sperm (Neuman et al., 2002). Supplementation of L-carnitine may also improve energy production from fatty acids to facilitate the hatching process in chicken embryos. The objective of the present study was to investigate whether reproductive efficiency would be improved by feeding L-carnitine to both parents over that of feeding L-carnitine to only the female or the male.
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MATERIALS AND METHODS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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From 0 to 3 wk of age, hatchlings were cage reared in a room of the Grower Research Unit of the Purdue University Poultry Research Center, with a maximum of 30 birds/cage, resulting in a density of 124 cm2/bird. At 3 wk of age, the sexes were separated and reassigned to cages in the same room, with 8 males and 11 to 12 females/cage and providing minimum floor space allowances of 464 cm2/male and 310 cm2/female. Each growing cage was equipped with 2 nipple drinkers, with feeder space allocations of 7.62 cm/male and 5.08 cm/female. Pullets and cockerels were maintained at these densities until cage transfer at 17 wk of age. Upon transfer, birds were maintained in the same room until termination of the experiment. Pullets were housed in a 61 x 36-cm-deep cage with 4 hens per cage, providing 549 cm2/bird. Cockerels were housed singularly in a 30 x 36-cm-deep cage, resulting in 1,080 cm2/bird. Feeder space for hens and roosters was 15 and 30 cm, respectively. There were 3 nipple drinkers per female cage and 2 per male cage. Mortality was recorded daily.
Room temperature was maintained at approximately 35°C the first week of the chick’s life. Ambient temperature was reduced 1 to 3°C each week until a temperature of 21°C was reached. Temperature was maintained at approximately 21°C and ranged from 18 to 24°C until termination of the experiment. Both the Grower and Layer Research Units were artificially ventilated with a negative-pressure system; evaporative cooling pads were used during hot weather.
Birds were vaccinated for Marek’s disease at hatch; Newcastle disease and infectious bronchitis at 2, 8, and 16 wk of age; infectious bursal disease at 3, 10, and 14 wk of age; and avian encephalomyelitis at 12 wk of age.
Continuous lighting at 10 lx was provided for the first 2 d posthatch. Light intensity was reduced to 5 lx at 3 d of age and maintained at that level for the remainder of the study. Fluorescent service lights were used periodically during regular light hours to assess bird health as well as to conduct artificial insemination. Light was reduced to 15 h/d from 3 d to 3 wk of age. From 3 to 17 wk of age, a constant 11-h photoperiod was maintained. At 17 wk of age, light hours were increased by 1 to 12 h of light. At 18 wk of age, light hours were again increased by 1 to 13 h of light. By 19 wk of age, 15 min/wk was added to the photoperiod until 16 h of light was achieved. The 16 h of light was maintained until termination of the experiment.
Birds were fed a corn- and soybean-based diet ad libitum. Diets were formulated to contain either 0 or 125 ppm of L-carnitine (Carniking, Lonza Inc., Allendale, NJ) beginning at hatch until 37 wk of age (Tables 1
and 2). The justification for feeding 125 ppm of L-carnitine beginning at 1 d of age in the current study was that this was the lowest dose that our laboratory had used to date in White Leghorns that had resulted in a sustained persistent increase in sperm concentration (Zhai et al., 2007). A red or green microtracer (Microtracers, San Francisco, CA) with no nutritive value was assigned by the feed mill manager to either the control (0 ppm) or treated (125 ppm of L-carnitine) diet; its purpose was to serve as a marker to ensure that the correct feed was placed in the appropriate feed trough. The investigators were blinded to the color code treatment assignment until termination of the experiment. Individual BW were determined at hatch before consumption of dietary treatments (designated as wk 0) and at 3 and 17 wk of age.
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From the 42 experimental units of semen, 6 semen pools were made, representing roosters from each of the 3 treatments of in ovo injections (saline, or 1 or 2 µmol of L-carnitine) and the 2 levels of dietary L-carnitine (0 and 125 ppm). Sperm concentration was determined for each of the 6 semen pools by hemocytometer counts (Bakst and Cecil, 1997). Within 30 min of collection, semen was inseminated into hens. A 2 x 2 factorial arrangement was used in which semen from roosters consuming either a control or L-carnitine-supplemented diet was inseminated into hens consuming either a control or L-carnitine-supplemented diet. Each of the 6 semen pools was used to inseminate 48 hens (12 replications/treatment of 4 hens each). Each of the 288 hens (48 hens/pool x 6 pools of semen) was artificially inseminated with 23.5 µL of semen at weekly intervals. Because roosters supplemented with dietary L-carnitine had higher sperm concentrations, this resulted in a mean insemination dose of 1.2 and 1.1 x 108 sperm/hen for L-carnitine and control hens, respectively.
Numbers of eggs laid per cage were recorded daily and hen-day egg production was calculated monthly. Egg and shell traits were determined every 4 wk when the breeding flock was 23, 27, 31, and 35 wk of age. For each of the 4 collection dates, 4 eggs were collected from each cage (experimental unit), resulting in a total of 288 eggs (4 eggs/cage x 3 in ovo injections x 2 diets x 2 rows/diet x 6 replications/row). Individual egg weights were determined. To obtain the weights of individual egg components, egg contents were separated by hand, and yolks were rolled on a wet paper towel to remove excess albumen and chalazae and then weighed. Percentage of yolk was calculated by dividing yolk weight by initial egg weight and multiplying the quotient by 100. The shells were rinsed with tap water, dried overnight at 60°C, cooled, and weighed. Shell weights included shell membrane weights. Percentage of shell was calculated by dividing shell weight by initial egg weight and multiplying the quotient by 100. Albumen weight was calculated by subtracting the yolk and shell weights from the respective egg weight; its proportional weight was calculated as described for percentage of yolk. Shell thickness, which included shell membrane thickness, was measured with a caliper (B. C. Ames Co., Waltham, MA) at 8 representative areas of each shell and then averaged. Specifically, 2 shell thickness measurements were made at the blunt end of the shell, 2 at the pointed end, and 4 near the equator.
One week before setting eggs for incubation, hatching eggs were marked by cage number, collected daily, and stored in a cooler at 10°C. Eggs were incubated in a James-way 252 incubator (James Manufacturing Co., Fort Atkinson, WI) and standard incubation practices were used relative to temperature, humidity, and egg turning. Hatch rates and hatchling group BW were determined at 4-wk intervals when the breeding flock was 30, 34, and 38 wk of age, resulting in 3 hatches. For each of the 3 hatches, 70 eggs were used from each of 12 treatments (3 in ovo injections x 2 female diets x 2 male diets), with 7 replicates or incubator tray levels per treatment. An experimental unit consisted of 10 hatching eggs. Eggs were transferred to hatching baskets on d 18 of incubation by placing 10 eggs in their own individual hatching compartment so as to keep experimental units separated. Because there were 7 d of hatching egg collection and storage in a cooler, eggs from different days of lay were evenly distributed to each of the 7 replicates and treatments. At 22 d of incubation, the number of chicks that hatched was counted for each experimental unit (10 hatching eggs) and group BW was determined. All unhatched eggs were opened to determine the number of infertile, early, mid, and late dead eggs. Hatched chicks were killed. Yolk sacs were collected and weighed from individually weighed hatchlings whose parents consumed either control or L-carnitine-supplemented diets and were injected in ovo with saline as embryos. Proportional yolk sac weights were calculated by dividing hatchling yolk sac weight by hatchling BW and multiplying by 100. Two yolk sacs from each experimental unit were chosen randomly for the determination of yolk sac lipids (AOAC, 2005) and fatty acid composition. Yolk sac lipids were extracted with chloroform:methanol (2:1, vol/vol). Lipids were then saponified and fatty acid methyl esters were prepared by transesterification with boron trifluoride in methanol (14%, wt/wt). Lipids extracted from yolk sacs were methylated (sodium methoxide) as follows: each sample was dissolved in dry toluene (1 mL) in a test tube with a Teflon-lined screw cap, 0.5 M sodium methoxide in anhydrous methanol (2 mL) was added, the solution was maintained at 50°C for 10 min, and glacial acetic acid (0.1 mL) was added, followed by deionized water (5 mL). The fatty acid methyl esters were extracted into hexane (2 mL), dried over anhydrous sodium sulfate, and filtered. Samples were analyzed by using a Varian 3900 gas-liquid chromatograph equipped with a flame-ionization detector with an 8400 autosampler, and a wall-coated open tubular fused-silica capillary column (30 m x 0.32 mm, Varian Inc., Walnut Creek, CA). Initial oven temperature was set at 175°C and held for 4 min. The temperature was increased by 3°C per min to a final temperature of 240°C, which was held for 30 min. All samples were introduced by split injection (1:33) and identified by comparison of their retention times with authentic standards. Fatty acid values are presented as area percentages (Matreya Inc., Pleasant Gap, PA). The saturation of the fat in the yolks was measured in 2 different ways. The iodine value was calculated as (% C16:1 x 0.950) + (% C18:1 x 0.860) + (% C18:2 x 1.732) + (% C18:3 x 2.616) + (% C20:1 x 0.785) + (% C22:1 x 0.731) (AOAC, 2005). The ratio of saturated fat to unsaturated fat was calculated as (16:0 + 18:0 + 20:0 + 21:0 + 22:0 + 24:0)/(16:1 + 16:2 + 16:3 + 18:1 + 18:2 + 18:3 + 20:1 + 20:3 + 20:4 + 20:5 + 22:4 + 22:6).
L-Carnitine analysis was performed on feed and egg yolks. For feed samples, 4 replicates each of the control and L-carnitine-supplemented breeder diets were collected for analysis of L-carnitine. Within each of the 4 dietary treatments (2 x 2 factorial arrangement of hens and roosters consuming 0 or 125 ppm of L-carnitine), 3 egg yolks were gently mixed from a hen injected in ovo with saline, 1 µmol of L-carnitine, or 2 µmol of L-carnitine as embryos, to form 4 pools of yolk. The procedure was repeated for a second replicate. Two replicates of the 4 pools of yolk samples, for a total of 8 yolk samples, as well as the 8 feed samples were sent to Metabolic Analysis Labs Inc. (Madison, WI) for measurement of L-carnitine. Samples were quantified in a neutralized perchloric acid extract by using a radioisotopic enzymatic method (Parvin and Pande, 1977). This experiment was conducted under guidelines approved by the Purdue University Animal Care and Use Committee.
Data were analyzed by ANOVA with repeated measurements (Steel et al., 1997) by using the mixed model procedure of the SAS system (SAS Institute, 2003). Dietary treatment (0 and 125 ppm of L-carnitine), the in ovo injection treatment, age, and sex (BW analysis only) were considered fixed effects. All interactions among main effects were tested in the statistical model. A 2 x 2 factorial arrangement was used for hatch of set eggs, hatch of fertile eggs, embryonic mortality, progeny BW at hatch, and yolk L-carnitine concentration. It consisted of 1) both parents consuming the control diet, 2) both parents consuming the L-carnitine-supplemented diets, 3) hens consuming the L-carnitine-supplemented diet and males consuming the control diet, and 4) hens consuming the control diet and males consuming the L-carnitine-supplemented diet. Tray of the incubator was used as a block for the incubation data. Experimental units for each data set are defined in the tables and figures. Differences of least squares means (Tukey-Kramer) were used to detect differences among means. For BW, a separate ANOVA was conducted at each age (0, 3, and 17 wk of age) because of differences in sample size. As birds aged, the population was reduced to meet the number of observations needed by breeding age. Because the BW at hatch (0 wk) was different between dietary treatments (P 
0.0001), an analysis of covariance with repeated measures was conducted by using hatchling BW as a covariate. Only birds that had BW data for all 3 ages (0, 3, and 17 wk of age) were used in the analysis of covariance. A t-test was conducted to determine whether the analyzed values of L-carnitine in feed were different from calculated values (Steel et al., 1997).
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RESULTS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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At hatch, chicks assigned randomly to the L-carnitine-supplemented diet had day-old BW that were smaller than the control chicks (P 0.0001), but by 3 wk of age, differences in BW between the 2 diets dissipated (Table 3). By 17 wk of age, the L-carnitine-supplemented birds had lower BW than controls (P 0.0001; Table 3). When using day-old BW as a covariate, the BW at 17 wk of age was still lower for birds consuming L-carnitine as compared with control birds (P 0.01), with no differences between diets at 3 wk of age (P = 0.99).
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). As expected, the female diet and not the male diet caused an increase in the concentration of yolk L-carnitine. Samples of egg yolks had increased L-carnitine per gram of wet yolk (P = 0.001) in hens consuming L-carnitine as compared with control hens (Figure 1). The interaction between male and female diets was not significant, indicating that the response was the same for both groups of females, regardless of whether they received semen from control or L-carnitine-fed roosters. Hatch of set eggs, hatch of fertile eggs, infertility, and embryonic mortality were unaffected by dietary L-carnitine and in ovo injection of L-carnitine, but significant interactions between male and female diets or with in ovo injections did occur with some of the hatch traits (Table 5). Further interpretation of significant interaction effects on hatch traits showed no consistent trends that were of biological interest.
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0.005; Table 5
). Hatchlings from breeder parents injected as embryos with 1 or 2 µmol of L-carnitine had greater BW than hatchings from breeder parents injected as embryos with saline (P = 0.001; Table 5). Absolute yolk sac weight, proportional yolk sac weight, and yolk sac lipid content were lower (P = 0.0001, P = 0.001, and P = 0.01, respectively) in hatchlings whose male and female parents both consumed the L-carnitine-supplemented diet as compared with parents that both consumed the control diet (Figure 2). Yolk weight and its lipid content were unaffected by the age of the parent, and the interaction of dietary L-carnitine in the parental diet with parental age was also NS (data not presented in tables or figures). In addition to the lowered yolk sac lipid content of hatchlings whose parents consumed L-carnitine (Figure 2), the proportions of fatty acids were altered (Table 6). Hens consuming L-carnitine produced hatched chicks with higher levels of yolk sac myristic (C14:0) and oleic (C18:1n-9) acids compared with yolk sacs of chicks hatched from control hens. The opposite trend was observed in yolk sac linoleic (C18:2n-6) and -linolenic (C18:3n-3) acids in that samples from control chicks were higher when compared with those collected from L-carni-tine chicks. In general, the fatty acid composition of yolks of older hens (38 wk) was altered when compared with hens of the earlier ages of 30 and 34 wk. More specifically, the percentages of myristic (C14:0), palmitic (C16:0), stearic (C18:0), oleic (C18:1n-9 and C18:1n-7), linoleic (C18:2n-6), conjugated linoleic (C18:2 cis-9, trans-11), and -linolenic (C18:3n-3) acids and calculated iodine values were lower in yolks of hatchlings from 38-wk-old hens compared with yolks of chicks from either 30- or 34-wk-old hens. Conversely, the percentage of cis-7,10,13,16-docosatetraenoic (C22:4n-6) acid and the saturated-to-unsaturated fatty acid ratio were higher in yolks from 38-wk-old hens compared with either 30- or 34-wk-old hens. Yolk sac arachidonic (C20:4n-6) acid was also higher in the chicks hatched from older hens of 38 wk of age as compared with younger hens of 30 wk, but not 34 wk of age. There were no diet x age interactions for yolk fatty acid composition.
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DISCUSSION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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), most likely because of the high numbers of sperm that were inseminated in the L-carnitine-fed and control hens (1.2 and 1.1 x 108 sperm/hen, respectively). If lower sperm numbers had been inseminated into hens (less than the optimal dose of 1 x 108 sperm/hen), beneficial results relative to fertility and hatch rate may have been obtained.
Rabie et al. (1997) reported that supplementation of 50, 100, or 500 ppm of dietary L-carnitine did not affect egg production, egg weight, shell weight, and shell thickness during the late laying period from 65 to 73 wk in a Hungarian brown hybrid line. However, all dietary levels of L-carnitine improved egg-white percentage as compared with the control diet, perhaps because of the higher metabolic rate in the magnum, where albumen is synthesized, and in the shell gland, where water is added to albumen during egg formation. These investigators also reported a lower yolk percentage in eggs of hens consuming dietary levels of L-carnitine. Celik et al. (2004) reported that egg weight, yolk weight, shell weight, and shell thickness were not affected by supplementation of 50 ppm of L-carnitine in the drinking water of 47-wk-old laying hens for 8 wk. In addition, proportional albumen weight was increased by L-carnitine supplementation. Yalcin et al. (2005) found that the addition of L-carnitine to laying quail diets did not affect egg production, egg shell thickness, and the percentages of egg shell, albumen, and yolk, but did increase egg weight. In our current experiment, supplementation of L-carnitine to hen diets starting at hatch did not affect egg production, egg weight, yolk weight, shell weight, and shell thickness (Table 4).
The higher concentrations of L-carnitine in the yolk of fresh eggs obtained from hens consuming 125 ppm of L-carnitine as compared with control hens (Figure 1) may have encouraged the utilization of fat by developing embryos, as indicated by the smaller yolk sac weights and lower yolk sac lipid content (Figure 2) in our present experiment. During embryonic development, approximately 50% of the initial yolk lipid is oxidized for energy production; the other 50% is incorporated into the body tissue and residual yolk of hatchlings (Lin et al., 1991). L-Carnitine supplementation of hen diets improved embryo yolk lipid mobilization. The mobilized yolk lipid may be used to produce energy or be incorporated into body tissues. However, broiler breeder hens fed lower levels of L-carnitine (25 ppm) for shorter durations (from 21 to 40 wk of age) had hatch rates, 18-d-old embryo yolk sac weights (expressed proportional to egg weight), and total yolk lipids similar to control hens (Peebles et al., 2007). The lower levels of L-carnitine fed to the feed-restricted broiler breeders (25 ppm), as compared with the current study in which White Leghorns consumed 125 ppm ad libitum, may have not been high enough to facilitate yolk lipid utilization during embryogenesis and subsequent hatch, although the investigators reported a 49% increase in total L-carnitine of yolks of fresh eggs obtained from breeders consuming 25 ppm of L-carnitine as compared with control hens (Peebles et al., 2007). Another distinct difference between the studies in addition to dosage of L-carnitine and type of chicken is that in the current study, the L-carnitine-supplemented diet was initiated at hatch, whereas in the study by Peebles et al. (2007) L-carnitine feeding began at 21 wk of age. In addition, our measurements for yolk sac weight were done at hatch, whereas in the study by Peebles et al. (2007), yolk sac weight was measured in 18-d-old embryos.
Not only was the lipid content lowered in the yolk of chicks hatched from hens fed L-carnitine (Figure 2), but the yolk fatty acid composition was altered as well (Table 6), suggesting that L-carnitine mediated changes in lipid metabolism. The presence of L-carnitine in the yolk may have caused the developing embryo and hatchling to selectively utilize a greater proportion of linoleic (C18:2n-6) and -linolenic (C18:3n-3) acids for growth and development over myristic (C14:0) and oleic (C18:1n-9) acids (Table 6), perhaps increasing their transfer efficiency out of the yolk to the embryo. An additional possibility is that the presence of carnitine in the yolk may have affected the efficiency of enzymes involved in fatty acid metabolism. Although the results of the current study showed no dietary effect on palmitoleic acid (C16:1n-7), Peebles et al. (2007) reported a reduced percentage of this particular fatty acid in the yolk sac of 18-d-old embryos derived from 27-wk-old broiler breeder hens consuming 25 ppm of L-carnitine, but this compositional change did not persist in 32- and 38-wk-old hens.
It is well known that hen diet can affect the fatty acid profile of the yolk (Cruickshank, 1934; Sell et al., 1968; Sim et al., 1973; Hargis and Van Elswyk, 1993); however, the relative change in yolk composition as a function of hen age is less well known, except in the case of broiler breeders (Latour et al., 1996, 1998; Burnham et al., 2001; Peebles et al., 2007). Nielsen (1998) reported that the yolks of egg-laying strains of chickens had an overall percentage of polyunsaturated fatty acids that was 20 to 25% higher in yolks from younger hens (21 wk of age) compared with older hens (57 wk of age), suggesting that egg yolks become more saturated as hens age. The present study agrees with the concept that eggs become more saturated as a function of hen age, as indicated by the increase in the overall ratio of saturated to unsaturated fatty acids and the decrease in iodine value.
Broiler breeder females fed 50 and 100 ppm of dietary L-carnitine had increased yolk L-carnitine concentrations and improved hatch rates of 4 and 3%, respectively, when compared with controls, although no indication was given of statistical significance among treatments (Leibetseder, 1995). In the current experiment, we found that supplementation of dietary L-carnitine as compared with the control diet increased yolk L-carnitine concentration (Figure 1), decreased yolk sac weight and yolk sac lipid content (Figure 2), and altered fatty acid composition (Table 6), but did not result in a subsequent improvement in hatchability and fertility (Table 5). Peebles et al. (2007) also reported no effect on hatch rate of eggs from broiler breeder hens consuming 25 ppm of L-carnitine as compared with control hens.
Typically, male and female layer breeders are raised together during grow-out, and they mate naturally during the breeding phase. Male breeders consume the same diets as female breeders. Our previous research showed that feeding L-carnitine to male breeders beginning at hatch increases sperm concentration (Zhai et al., 2007). Female layer breeders consuming the same diets as males that are supplemented with L-carnitine can do so without causing harm to the number and quality of hatching eggs.
The lower hatchling BW of progeny from roosters consuming L-carnitine as compared with progeny hatched from control roosters (Table 5) and the significant in ovo injection effects on egg weight, albumen weight, shell weight (Table 4), and progeny BW (Table 5) were unexpected and the causes are unknown. The breeder parents were given the in ovo feeding of L-carnitine at 18 d of incubation, so it is perplexing that this effect persisted into adulthood, with subsequent effects on egg component weights and progeny BW. For the dietary male effect attributable to L-carnitine supplementation on hatchling progeny BW, a remote possibility could have been genetically derived. Roosters who later served as parents in the current study were assigned randomly to 0 and 125 ppm at hatch; the hatchlings assigned to the L-carnitine-supplemented diets were smaller than the control hatchlings before the initiation of treatments (P 0.0001; Table 3).
We concluded that supplementation of diets with L-carnitine did not result in a subsequent improvement in hatch rate, most likely because of the high numbers of sperm inseminated artificially to both control and L-carnitine-supplemented hens. Higher concentrations of L-carnitine in the yolk of fresh eggs obtained from hens consuming L-carnitine as compared with control hens may have encouraged the utilization of fat by developing embryos, as indicated by the smaller yolk sac weights and lower yolk sac lipid content. However, if more yolk fat was indeed used by the developing embryo whose female parent consumed L-carnitine, it did not culminate in improved hatchability. Additional studies will be needed to determine whether carnitine-induced increases in yolk fat mobilization in hatchlings affect chick livability during the brooding period.
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ACKNOWLEDGMENTS |
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FOOTNOTES |
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Received for publication August 3, 2007. Accepted for publication February 10, 2008.
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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