Early feeding in the hatcher influences physiological indicators of nutritional status and hydration in broiler chicks during the immediate post-hatch period

Abstract

As early feeding (EF) is considered a promising strategy to improve broiler welfare, performance and health, new approaches to implement EF are becoming increasingly popular in the poultry industry. EF provided in the hatcher, commonly referred to as in-hatcher feeding, can offer a solution for the feed and water deprivation period experienced by chicks in traditional hatchery systems. However, in-hatcher feeding is still developing and scientific research regarding its impact on performance, health and welfare is limited. Therefore, the present study was designed to determine specific short-term physiological responses to EF in broiler chicks hatched in a hatcher, focusing on stress indicators and indicators for inferring time to first feed and water intake. A total of 320 Ross 308 broiler chicks, housed in 32 hatching baskets (10 chicks per basket; 8 replicates per treatment), were randomly assigned to one of four post-hatch feeding treatments until 48 h post-hatch: immediate access to feed and water (FW), to feed only (F), to water only (W), or deprived of both feed and water (D). At 4, 14, 24, 36, and 48 h post-hatch, blood and tissue samples (proventriculus and residual yolk) were collected to assess chick body weight (BW), residual yolk weight, plasma osmolality, plasma arginine vasotocin (AVT), plasma corticosterone (CORT) and ghrelin mRNA. Data were analysed using mixed models with the hatching basket as the experimental unit. From 24 h - 48 h post-hatch, BW differed between treatments with FW consistently showing the highest BW when compared to the other treatments (p < 0.001). An age-related increase in plasma osmolality was observed in water deprived treatments (D and F) from 24 - 48 h post-hatch, while levels in W and FW decreased slightly (p < 0.001). In D, AVT levels increased sharply from 24 h post-hatch onwards, whereas FW consistently showed lower levels compared to D (p = 0.005). Similarly, plasma CORT concentrations were lower in FW compared to D (p = 0.010) at 36 and 48 h post-hatch. Ghrelin mRNA expression did not differ between treatments during the first 14 h post-hatch but was consistently lower in FW when compared to D, F and W (p < 0.001) thereafter. Conversely, no treatment effects were observed for residual yolk weight. In conclusion, the physiological and neuroendocrine responses observed between 14 and 24 h post-hatch suggest a transition in metabolic and hydration status that is consistent with the onset of feeding and drinking during this time window. Additionally, higher body weights and reductions in corticosterone levels were only observed when both feed and water were provided.

Introduction

According to the Food and Agriculture Organisation (FAO), six billion broiler chicks are reared annually in the European Union (FAO, 2024). The vast majority of these chicks hatch in conventional hatcheries, whereas chicks hatching on-farm are still underrepresented (De Jong and Gunnink, 2019; de Jong et al., 2020). In conventional hatcheries, newly hatched chicks typically do not have access to feed or water until placement in the broiler house (De Jong et al., 2017), and are therefore entirely dependent on nutrients provided by the residual yolk. This period of feed and water deprivation post-hatch may last up to 48 h and is largely caused by the biological hatching window (Powell et al., 2016; Willemsen et al., 2010) in combination with hatchery procedures and chick transportation times (De Jong et al., 2017). In this context, alternative strategies that provide feed and water immediately post-hatch, commonly referred to as early feeding (EF), are emerging as a promising approach in broiler production. Currently used EF strategies, including in-hatcher feeding (Willemsen et al., 2010), on farm hatching (de Jong et al., 2020) and in ovo feeding (Joanna et al., 2017), are relatively recent and their potential benefits are not yet fully understood. Previous research in EF have already shown benefits for intestinal development (Gaweł et al., 2025; Li et al., 2022) and overall production efficiency (De Jong et al., 2017; Li et al., 2022; Smets et al., 2026), yet evidence regarding the effects on short-term welfare, including physiological indicators of nutritional status and hydration, and time to first feed intake remains limited (De Jong et al., 2017). The EFSA report on the welfare of broilers on farm, identifies body weight loss and mortality as the most useful animal-based welfare indicators for evaluating short-term welfare in hatchery-hatched chicks (Nielsen et al., 2023). However, both indicators are low in sensitivity and specificity as increases appear only after prolonged deprivation and can be affected by multiple other factors, including disease or management procedures. Additional animal-based physiological indicators could offer better insight into the physiological consequences of delayed access to feed and water in commercial hatcheries and the potential benefits of EF.

Ghrelin is a peptide predominantly produced in the proventriculus and involved in the regulation of feeding-related signalling in birds (Kaiya et al., 2012). In six-day-old layer chicks, both plasma ghrelin concentrations and ghrelin mRNA expression in the proventriculus increased following 12 hours of fasting and returned to baseline levels after refeeding (Kaiya et al., 2007). In line with these findings, delayed feed access for 72 h in broiler chicks significantly upregulated ghrelin mRNA expression, resulting in 2.1-fold higher levels than those observed in fed controls (Yu et al., 2016). This response appeared to be rapidly modulated by refeeding, as ghrelin mRNA expression decreased sharply within 4 h after the onset of feeding, reaching levels that were significantly lower than those observed in fed controls (Yu et al., 2016). These findings suggest that ghrelin is responsive to short-term changes in nutritional status in birds and can therefore act as a physiological correlate of feed intake. In conventional practices, delayed access to water may challenge osmoregulatory homeostasis in newly hatched chicks. Blood plasma osmolality has been shown to be a valuable physiological marker of dehydration in 6-week-old broilers (Chaturvedi et al., 1997; Knowles et al., 1996; Vanderhasselt et al., 2013), but has not yet been studied in newly hatched chicks under commercial hatchery conditions. Complementary to blood osmolality, arginine vasotocin (AVT) is a central neuropeptide regulating water balance in birds (Goldstein, 2006), which increases during water deprivation (Chaturvedi et al., 1997; Seth et al., 2004). Monitoring these physiological indicators in broiler chicks provided with EF can therefore provide insight into short-term physiological responses associated with nutritional status and hydration. Moreover, temporal changes of these indicators in early-fed chicks may offer indirect information on the timing of first feed and water intake after hatch, which is essential for evaluating the relevance of EF strategies and could clarify the inconsistent results reported in previous studies on time to first feed intake. Boyner et al. (2021), stated that chicks initiated feeding only after 24 h post-hatch, whereas earlier literature reported that chicks consumed an average of 1.5 g of feed within the first 24 h (Pinchasov and Noy, 1993).

Studies investigating the effects of feed and water deprivation on the stress response in the immediate post-hatch period remain limited and inconclusive. Tong et al. (2015) for example, did not observe a direct relationship between the duration of the holding period and plasma corticosterone (CORT) levels, while van de Ven et al. (2013) and Wijnen et al. (2022) reported that EF reduces CORT levels immediately after hatching.

The present study aimed to investigate short-term physiological responses of broiler chicks hatched in a hatcher to immediate post-hatch access to feed and/or water. Specifically, it evaluated how providing feed, water, or both during the first 48 h post-hatch affects physiological indicators of nutritional status, hydration, and stress. By characterizing the temporal dynamics of these parameters, the study further examined the onset of feeding and drinking post-hatch in a hatchery context.

Materials and methods

Ethics statement

The trial was carried out in compliance with the European Directive 2010/63/EU on the protection of animals used for scientific purposes. Ethical approval was granted by the Ethics Committee of the Research Institute for Agriculture, Fisheries and Food (ILVO), Merelbeke-Melle, Belgium, under authorization 2024/461.

Study design

The experiment was conducted over the first 48 hours post-hatch and included four feeding treatments applied throughout and 8 replicates per treatment. Chicks in the feed and water (FW) treatment had immediate access to both feed and water after hatching. The feed-only (F) treatment received feed but no water, while the water-only (W) treatment received water but no feed. In the deprivation (D) treatment, chicks were withheld from both feed and water. A complete randomized block design was applied, using the hatching window as the blocking factor to ensure balanced representation of all treatments within each hatch window. The hatching basket, a plastic container in which newly hatched chicks remain until pull, was used as the experimental unit.

Animal management

A total of 650 Ross 308 18-day incubated eggs, originated from a 36-week old parent flock, were obtained from a commercial hatchery (Vervaeke, Tielt, Belgium) and further incubated in a hatcher (X-Streamer2HOX-H, Petersime, Belgium) at the ILVO research centre. For this experiment, only chicks hatching within the middle 60% of the hatch window were selected. During hatching, chicks were evenly and randomly distributed across the four treatments, ensuring a balanced allocation based on time of hatching. The chicks assigned to each treatment were placed in a second similar hatcher after hatch, whereas the non-selected chicks were used in a separate, unrelated study. Hatcher temperature was regulated based on eggshell temperature, which was maintained between 99 and 99.5°F during hatching. Chick rectal temperature was monitored every 8 h using a CVET thermometer (Covetrus, Portland, Maine, USA), and the chick holding hatcher settings were adjusted accordingly to maintain chicks body temperature between 39.5 – 40.5°C. Chicks were housed in groups of 10 in commercial hatching baskets with no bedding and exposed to continuous white LED strip lighting. Feed and water were provided ad libitum according to the treatment using separate open plastic troughs (30 cm × 5 cm). Water was refreshed every 8 h. A standard starter feed, offered as a mash, was used for the treatments receiving early feeding. The wheat, corn, soybean meal based diet was formulated as a standard broiler diet (Table 1) to meet nutritional requirements according to the Ross 308 nutrition specifications (Aviagen, 2022).

Table 1.

Diet formulation (% fresh matter) (A) and calculated nutrient composition of the diet (B) offered in the hatcher for the Ross 308 broiler chicks.

A
Wheat	46.57
Maize	15.00
Soybean meal	25.11
Soybeans toasted	5.00
Soy oil	1.00
Animal fat	3.04
Premix (vitamins & minerals)	1.00
Limestone	0.55
Bi-calcium phosphate	1.20
Sodium chloride (NaCl)	0.13
Sodium bicarbonate	0.37
DL-methionine	0.34
L-Lysine HCl	0.37
L-Threonine	0.17
Coccidiostaticum	0.05
NSP Enzyme	0.01
Fytase	0.10
B
DM (g/kg)	111
Crude protein (g/kg)	205
Crude fat (g/kg)	71.0
Crude fiber (g/kg)	29.5
Metabolizable energy (MJ/kg)	11.8
Dig. lysine (g/kg)	11.5
Ca (g/kg)	9.0
Available P (g/kg)	4.3

Data collection

Sampling

Two chicks per basket were randomly selected for sampling at 4, 14, 24, 36, and 48 h post-hatch, with post-hatch age corresponding to their exact time since leaving the eggshell. Prior to killing via decapitation, selected chicks were weighed using a high-precision scale (DM-11000, UWE, Taiwan; accuracy: 0.1 g). Blood was collected in K3EDTA tubes (13 × 75 mm, non-ridged Vacuette®, Greiner Bio-One, Kremsmünster, Austria) and immediately centrifuged at 3,000 x g for 10 minutes at 4 °C to separate plasma. Plasma was stored at 4 °C until analysis later that day. Following blood collection, the residual yolk was removed and weighed, the proventriculus was excised, washed in phosphate-buffered saline (PBS), and submerged in RNAlater® (Thermo Fisher Scientific, Waltham, MA, USA). Tissues were stored at –20 °C prior to RNA isolation and subsequent qPCR analysis for ghrelin mRNA expression.

Plasma analysis

Plasma osmolality was measured using the freezing point depression method with an Osmostation OM-6060 (CRI, Zwijnaarde, Belgium). A sample volume of 225 µL plasma was used for each measurement. Plasma arginine vasotocin concentrations were determined by enzyme-linked immunosorbent assay (ELISA) according to the manufacturers protocol (Chicken Arginine Vasopressin (AVP) ELISA Kit, BlueGene Biotech, China). Corticosterone levels in plasma were measured using an ultra-performance liquid chromatography-tandem mass spectrometry (UPLC-MS/MS) analysis method. Therefore, plasma samples (250 µl) were spiked with 25 µl of internal standard solution (CORT-d8, 100 ng/ml) and 25 µl of methanol (or CORT working solution in methanol for calibrators/QCs), vortex mixed, and subjected to a liquid-liquid extraction with 3 ml of diethyl ether. After 20 min of rolling on a roller mixer and subsequent centrifugation (3,000 rpm, 10 min, 4 °C), the organic phase was evaporated to dryness under nitrogen at 40 °C. The residue was reconstituted in 125 µl of methanol and 125 µl of water, filtered over a Nylon syringe filter (0.22 µm, 13 mm, Merck, Overijse, Belgium), and a 5 µl sample aliquot was injected into the UPLC-MS/MS system for quantification of corticosterone levels, as further detailed in (Soster et al., 2026).

RNA extraction

Total RNA was isolated from proventriculus and ileum samples using the Aurum Total RNA Mini Kit (Bio-Rad, CA, USA), with a modified initial homogenisation step. Briefly, tissue samples (35–45 mg) were homogenised in 500 µL TRIzol Reagent (Invitrogen, Waltham, MA, USA) using six 2.3 mm silica beads and one 3.2 mm chrome steel bead in a Qiagen TissueLyser II for 3 × 3 min at 30 Hz. After addition of 120 µL RNase-free chloroform, samples were centrifuged for 15 min at 14,000 × g at 4°C. The aqueous phase was mixed with 600 µL 60% ethanol and loaded onto Aurum RNA binding columns. Subsequent column purification, including DNase I treatment, washing, and elution in 30 µL elution solution, was performed according to the manufacturer’s instructions. RNA concentration was measured at 260 nm using a NanoDrop® ND-1000 spectrophotometer (Thermo Fisher Scientific, Waltham, MA, USA). Purified RNA was further treated using the TURBO DNA-free™ Kit (Invitrogen, Waltham, MA, USA). Total purified RNA (1 µg) was reverse transcribed to cDNA using the iScript cDNA Synthesis Kit (Bio-Rad) and stored at –20°C until further use.

Primer design and qRT-PCR analysis

The primer sequences used for the target gene ghrelin (Ghazanfari et al., 2010), as well for the housekeeping genes TATA-box Binding Protein (TBP), Hydroxymethylbilane Synthase (HMBS) and β-Glucuronidase (GUSB) (Chen et al., 2023), were obtained from published literature. Primer sequences were verified by confirming their presence within the corresponding gene sequences using the NCBI’s Nucleotide database (https://www.ncbi.nlm.nih.gov/nucleotide/). Primer efficiency was evaluated using serial dilutions of a cDNA sample pool (1, 1:2, 1:4, 1:8; 1:16) to obtain the amplicon efficiency (E), correlation coefficients (R²) and slope. The melting curves were analysed in Bio-Rad CFX Maestro, and for all primer pairs, a single peak was detected. Gene names, accession numbers, primer sequences, E, R² and slope are given in Table 2. After confirmation of the specificity and efficiency of every primer set, Real-time quantitative PCR reactions were run in triplicate. The 12 µl reaction mixture consisted of: 6 µl iQ SYBR Green Supermix (Bio-Rad); 4 µl HPLC-grade water (Merck Millipore, Overijse, Belgium) containing 0.5 µM each of the forward and reverse primers; 2 µl undiluted cDNA for ghrelin. The PCR protocol (40 cycles) was performed on a CFX384 Touch Real-Time PCR System with a C1000 Thermal Cycler (Bio-Rad). The protocol used was 95°C for 20 s followed by 60°C for 30 s and 72°C for 30 s repeated 40 times. The results were analysed by using the Bio-Rad CFX Maestro. Quantification cycle (Cq) values were obtained using auto baseline settings, and these were applied per primer set. The obtained Cq values were then imported in QBase+ for fully automated analysis and interpretation of reference gene stability using the GeNorm algorithm (Hellemans et al., 2008). Three reference genes were used as internal controls for each gene of interest. The stability (M-values) of the individual reference genes of all samples is given in Table 2. Relative quantification was performed automatically using qbase+ software. The resulting calibrated normalized relative quantification (CNRQ) values were scaled to the mean and log-transformed prior to statistical analysis.

Table 2.

Primer sequences used in the real-time RT-PCR analysis.

Gene	Accession number		Sequence (5-3′)	E (%)	R²	Slope	M-value
Ghrelin		AY303688.1	5′-CCT TGG GAC AGA AAC TGC TC-3′ 5′-CAC CAA TTT CAA AAG GAA CG-3′	110	0.988	−3.103	-
HMBS1		XM_046932409.1	5′-GGC TGG GAG AAT CGC ATA GG-3′ 5′-TCC TGC AGG GCA GAT ACC AT-3′	106.7	0.978	−3.172	1.458
GUSB2		NM_001039316.2	5′-GGC AGA CTG GTC CTG TTG TTG-3′ 5′-GGG TCC TGA GTG ATG TCA TTG A-3′	90.6	0.981	−3.570	1.342
TBP3		D83135.2	5′-GCG TTT TGC TGC TGT TAT TAT GAG-3′ 5′-TCC TTG CTG CCA GTC TGG AC-3′	98.8	0.990	−3.351	1.636

¹Hydroxymethylbilane Synthase.

²β-Glucuronidase.

³TATA-box Binding Protein.

Statistical analysis

Statistical analyses were performed using SAS 9.4 through SAS Studio via SAS OnDemand for Academics (SAS Institute Inc., Cary, NC, USA). Since repeated measurements within baskets were assumed to be correlated, the data were analysed using a mixed model with replicate and the hatching basket as random effects, and treatment, sex, age and the two-way interactions treatment-age and treatment-sex as fixed effects. To account for potential effects of hydration status on plasma AVT and corticosterone concentrations, plasma osmolality was included as a continuous covariate in the mixed model. Pairwise comparisons between treatments were performed using the post-hoc Tukey’s test to obtain adjusted p-values. Statistical significance was set at p < 0.05.

Results

Body weight and residual yolk weight

A strong age × treatment interaction (p < 0.001) indicated that treatment effects varied across post-hatch age for BW. Post-hoc comparisons across age revealed significant differences between D and W (p = 0.006), D and FW (p < 0.001), F and W (p = 0.031), F and FW (p < 0.001) and W and FW (p = 0.034) (Table 3). Estimates indicated that BW was comparable among treatments until 14 h post-hatch but diverged from 24 h onward, with increasing differences up to 36 and 48 h post-hatch. Treatments with access to water (FW and W) showed stable BW until 24 h post-hatch but diverged afterwards. By 36 and 48 hours post-hatch, chicks in FW showed a gradual increase in BW, whereas chicks in W lost weight. F and D gradually lost weight over time (Fig. 1). For absolute residual yolk weight, no significant effects of treatment or sex were observed (Fig. 2).

Table 3.

Effect of post-hatch availability of feed and/or water on body weight, yolk weight, plasma osmolality, AVT, CORT and ghrelin mRNA expression. Data are presented as least-squares means ± standard error. Means with different superscript differ significantly (P < 0.05) within a same row.

	FW	F	W	D	P value
Body weight (g)	45.57a ± 0.49	41.33c ± 0.51	43.49b ± 0.50	40.88c ± 0.49	<0.001
Yolk weight (g)	4.08 ± 0.10	4.00 ± 0.11	3.87 ± 0.11	3.88 ± 0.10	0.437
Plasma osmolality (mOsm/kg)	323.88b ± 0.93	343.98a ± 1.03	322.88b ± 0.98	343.22a ± 0.92	<0.001
AVT (pg/mL)	350.2b ± 30.0	406.6ab ± 32.35	448.4a ± 31.84	482.5a ± 25.56	0.010
CORT (ng/mL)	8.25b ± 2.11	11.63ab ± 2.04	15.07a ± 2.03	13.70ab ± 2.0	0.011
Ghrelin mRNA expression (log)	−0.50b ± 0.06	0.12a ± 0.07	0.18a ± 0.06	0.22a ± 0.07	<0.001

FW = feed and water. F = feed only. W = water only. D = deprived.

Fig. 1.

Effect of post-hatch availability of feed and/or water on body weight at 4, 14, 24, 36, and 48 hours post-hatch. Data are presented as least-squares means ± standard error.

Fig. 2.

Effect of post-hatch availability of feed and/or water on residual yolk weight at 4, 14, 24, 36, and 48 hours post-hatch. Data are presented as least-squares means ± standard error.

Plasma osmolality

A strong age × treatment interaction (p < 0.001) indicated that treatment effects varied across post-hatch age. Post-hoc comparisons across age revealed significant differences between D and W (p < 0.001), D and FW (p < 0.001), F and W (p < 0.001), and F and FW (p < 0.001) (Table 3). Estimates indicated that plasma osmolality was comparable among treatments at 4 h post-hatch but diverged from 14 h onward, with increasing differences up to 48 h post-hatch. Treatments with access to water (FW and W) showed significantly lower osmolality levels compared with those deprived of water (D and F) (Fig. 3).

Fig. 3.

Effect of post-hatch availability of feed and/or water on plasma osmolality at 4, 14, 24, 36, and 48 hours post-hatch. Data are presented as least-squares means ± standard error.

Plasma arginine vasotocin

A strong age × treatment interaction (p < 0.001) indicated that treatment effects varied across post-hatch age. Post-hoc comparisons across age only revealed a difference between D and FW (p = 0.048), and between W and FW (p = 0.048) (Table 3). Estimates indicated that AVT levels were comparable among treatments at 4 and 24 h post-hatch, but treatment differences became more apparent from 36 h onwards. At 36 h, D showed the highest AVT levels, whereas F and FW treatments remained lower. By 48 h, AVT concentrations increased in W, F and FW but declined in D (Fig. 4).

Fig. 4.

Effect of post-hatch availability of feed and/or water on AVT at 4, 14, 24, 36, and 48 hours post-hatch. Data are presented as least-squares means ± standard error.

Plasma corticosterone

A significant effect of age (p < 0.001) and treatment (p = 0.004) was observed. Post-hoc comparisons across treatments showed that CORT levels differed significantly between W and FW (p = 0.008) (Table 3). Estimates indicated that CORT levels were lowest in the FW treatment compared to the other treatments (Fig. 5).

Fig. 5.

Effect of post-hatch availability of feed and/or water on CORT at 36, and 48 ho urs post-hatch. Data are presented as least-squares means ± standard error.

mRNA expression

A strong age × treatment interaction (p < 0.001) indicated that treatment effects differed across post-hatch ages. Post-hoc comparisons across treatments revealed significant differences between FW and W (p < 0.001), FW and F (p < 0.001), and FW and D (p < 0.001), whereas F, W and D were not significantly different from each other (Table 3). Ghrelin mRNA expression was comparable among treatments up to 14 h post-hatch, after which the treatments started to diverge. Starting from 24 h post-hatch onwards, the FW treatment consistently showed lower ghrelin mRNA expression levels compared to the other treatments (Fig. 6).

Fig. 6.

Effect of post-hatch availability of feed and/or water on mRNA ghrelin expression at 4, 14, 24, 36, and 48 hours post-hatch. Data are presented as least-squares means ± standard error.

Discussion

Early feeding provided in the hatcher led to measurable changes in physiological parameters post-hatch, suggesting that the majority of chicks initiate feeding and drinking between 14 and 24 h post-hatch. Our findings further indicate that water must be available to support feed intake, and that only the combination of both feed and water supports growth and reduces CORT levels. EF did not significantly affect residual yolk weight.

Access to feed and water in the immediate post-hatch period influences early BW development in broiler chicks. In conventional hatchery settings, when no EF is provided, chicks lose approximately 8% of their BW within the first 24 hours (Careghi et al., 2005a; Gonzales et al., 2003; Noy and Sklan, 1999), increasing to 17% at 48 hours post-hatch (De Jong et al., 2017). Consistent with this, D and F chicks in our study exhibited comparable weight loss, with BW losses of 8% (D) and 11% (F) at 24 hours post-hatch, increasing to 15% (D) and 16% (F) at 48 hours post-hatch. This early BW loss is generally attributed to utilisation of the residual yolk, dehydration and catabolism of the pectoralis muscle (Careghi et al., 2005; Maman et al., 2019; Pinchasov and Noy, 1993). The less pronounced BW loss observed in W chicks suggests that dehydration contributed substantially to post-hatch BW loss in the present study. This aligns with findings of Incharoen et al. (2015) who reported reduced BW loss in chicks provided with an aqua agar containing 95% of water and 5% of agar powder during 24 h of post-hatch transport. Together, these findings highlight the mitigating effect of water on post-hatch BW loss. Conversely, providing feed alone did not reduce weight loss during the observation period, which is likely related to inhibited feed intake in the absence of water. Studies in older broilers have demonstrated that 24 or 48 h water deprivation markedly reduces feed intake by 21% and 48%, respectively, compared with non-deprived controls (Koike et al., 1983). This further supports the prandial link between feeding and drinking, a common phenomenon observed in poultry (Nielsen et al., 2010). Previous research on EF and body weight gain has shown that growth does not occur before 24 hours post-hatch (Noy and Sklan, 1999), which is in line with our observation that both feed and water must be available for more than 24 h to support growth. This early growth can be explained by the presence of functionally active digestive enzymes (Nitsan et al., 1991) and intestinal nutrient transporters (Mahagna and Nir, 1996) immediately after hatch, indicating that chicks are physiologically capable of processing early feed intake and using absorbed nutrients for body weight gain once both feed and water are available.

Absolute yolk weight gradually decreases with post-hatch age and is not affected by EF. It is known that the residual yolk is an important endogenous nutrient source during the post-hatch period and remains available to the chick until five days post-hatch (Lamot, 2017). It contains maternal antibodies (Ulmer-Franco et al., 2012) and valuable nutrients that support growth (Murakami et al., 1992; Turro-Vincent et al., 1994) and intestinal development (Noy and Sklan, 1999). Consequently, efficient utilization of the residual yolk is regarded as important for optimal chick health and performance. EF has been proposed to enhance post-hatch yolk resorption as increased intestinal activity in fed chicks may enhance the transport of yolk through the intestine (El - Husseiny et al., 2008; Noy et al., 1996). Bhanja et al. (2009) reported higher residual yolk uptake in early-fed chicks compared to 48 h deprived chicks. However, findings across studies are inconsistent as several reports have shown no effect of EF on yolk sac resorption during the first 3 days after hatching when chicks were subjected to a 36 – 72 h fasting period (Gonzales et al., 2003; van den Brand et al., 2010; van der Wagt et al., 2020). Our findings align with the latter observations and do not support the hypothesis that EF alters yolk sac utilization, suggesting that the residual yolk is resorbed at a relatively constant rate independent of exogenous feed intake.

Dehydration in broiler chicks during the post-hatch period is reflected by increased plasma osmolality. This is consistent with earlier research in 6-week-old broilers showing that 24 h of water deprivation is sufficient to detect differences in plasma osmolality (Nouwen et al., 1984; Vanderhasselt et al., 2013). Moreover, the gradual increase in D and F chicks observed in our study aligns with reports of rising plasma osmolality during prolonged water deprivation (Saito and Grossmann, 1998; Stallone and Braun, 1986; Wurtz et al., 2024). Elevated plasma osmolality is regarded as one of the most suitable physiological indicators of dehydration in adult broilers following 24 hours (Vanderhasselt et al., 2013) or 48 hours (Arad et al., 1985) of water deprivation. However, to the best of our knowledge, this is the first report of its application in newly hatched chicks during the post-hatch holding period. Dehydration in day-old chicks has previously been assessed based on alternative indicators, including increased plasma protein concentration and elevated haematocrit. Wyatt et al. (1986) reported elevated plasma protein concentration and haematocrit levels in chicks held in the incubator for an additional 30 h after hatching when compared to controls. In the present study, chicks with access to water showed physiological changes consistent with improved hydration before 24 h post-hatch. This pattern may indirectly reflect the onset of water intake during the first 24 h after hatching. However, because actual water intake was not directly quantified, this interpretation remains inferential and cannot be confirmed solely on the basis of the physiological parameters assessed in this study. In addition, plasma osmolality may not respond immediately after water intake, meaning that water intake could have started before changes in osmolality became detectable. Furthermore, the extent to which the observed increase in plasma osmolality corresponds to the experience of thirst remains unclear, as thirst perception is considered to be mediated by multiple sensory feedback mechanisms (Vanderhasselt et al., 2013). Cockram (2007) stated that thirst develops once physiological changes associated with the activation of homeostatic mechanisms to conserve water become apparent. Such physiological changes may therefore serve as early indicators of thirst-related responses in newly hatched chicks (Cockram, 2007). Arginine vasotocin (AVT) is a central neuropeptide involved in the regulation of water balance in birds (Goldstein, 2006). AVT levels are known to increase during water deprivation (Chaturvedi et al., 1997; Seth et al., 2004) and have been reported to correlate positively with changes in plasma osmolality (Arad and Skadhauge, 1984; Koike et al., 1983; Yahav et al., 2004). Previous studies have shown that water deprivation periods of 8 h (Saito and Grossmann, 1998), 24 h (Nouwen et al., 1984; Robinzon et al., 1990), or 48 h (Arad et al., 1985) were associated with increased AVT levels, which can rise 2- to 6-fold during dehydration (Chaturvedi et al., 2000). Physiologically, AVT acts to conserve body water by reducing glomerular filtration rate and by enhancing tubular water reabsorption in the kidneys (Skadhauge, 1981). As such, AVT release can be proposed as an early physiological indicator of thirst. In the present study, treatment-related differences in AVT levels only became apparent from 36 h post-hatch, while temporal patterns in AVT dynamics were also observed over the post-hatch period. In D, AVT levels increased progressively over time, reaching a plateau phase at 36 hours post-hatch. This pattern is generally consistent with previous reports describing a linear increase in AVT with increasing duration of water deprivation (Seth et al., 2004), although the plateau was reported to occur slightly later, after 48 hours (Stallone and Braun, 1986). Despite the overall trends, the expected positive correlation between plasma osmolality and AVT was not consistently observed in our study. Notably, W chicks showed a pronounced increase in AVT after 24 hours post-hatch while plasma osmolality levels even showed a mild decline at that time. In F chicks, AVT levels were similar to those in FW, although plasma osmolality was markedly higher. FW chicks exhibited a slight but consistent increase in AVT with post-hatch age, whereas plasma osmolality levels even showed a mild decline. These findings suggest that AVT release is not exclusively driven by changes in plasma osmolality and that additional regulatory factors may contribute to AVT dynamics during the early post-hatch period. Studies have shown that AVT may also be involved in activation of the HPA axis, as AVT concentrations have been reported to increase in response to acute stressors such as immobilisation (Aman et al., 2016) and feed deprivation (Kadhim et al., 2020). A possible link with the HPA-axis activation is further supported by the presence of AVT receptors on corticotropic cells (Cornett et al., 2013) and by studies showing that intracerebral AVT administration increases corticosterone concentrations (Masunari et al., 2016). However, the present results do not provide clear evidence for such a relationship, as temporal patterns of AVT and CORT were not consistent. Other described functions of AVT include roles in reproduction, oviposition, sexual differentiation, and sexually dimorphic behaviour (Jurkevich and Grossmann, 2003). However, these functions are less likely to explain AVT dynamics in the present study because of the young age of the chicks and the absence of sex-related differences. The mechanisms underlying the observed treatment- and age-related changes in AVT therefore warrant further investigation.

Although ghrelin concentrations generally increase during fasting and decrease after refeeding (Kaiya et al., 2007; Shousha et al., 2005), ghrelin appears to exert predominantly anorexigenic effects in chickens. This anorexigenic role is supported by studies showing that both central (Furuse et al., 2001; Saito et al., 2002) and peripheral (Geelissen et al., 2006; Shousha et al., 2005) ghrelin administration reduce feed intake in birds. This effect is thought to be mediated through activation of the adrenocorticotropic hormone (ACTH) pathway, as reductions in feed intake were attenuated when corticotropin-releasing hormone (CRH) antagonists were administered (Saito et al., 2005). However, it should be noted that exogenous ghrelin administration and endogenous ghrelin mRNA expression are not directly equivalent. Therefore, the functional role of endogenous ghrelin expression in relation to feed intake remains uncertain and warrants further investigation. Nevertheless, although a direct role as a hunger hormone in chickens remains unclear, there is consistent agreement that ghrelin concentrations respond to feed intake, as decreases in plasma ghrelin have been observed as early as 3 h (Shousha et al., 2005) and 6 h after refeeding (Kaiya et al., 2007). This suggests that ghrelin may serve as a useful indicator for assessing the timing of first feed intake during the post-hatch period. In the present study, ghrelin expression levels diverged from the other treatments at 24 h post-hatch in FW chicks, demonstrating that feeding-related physiological changes had occurred before this timepoint. Considering that decreases in ghrelin levels after refeeding have previously been observed within 3–6 h, this divergence may indicate that feed intake had already occurred before 24 h post-hatch, most likely during the 14–24 h post-hatch interval. However, actual feed intake was not directly quantified. This indirect estimate of the possible timing of first feed intake appears to contrast with findings from Boyner et al. (2021), who reported that only 5% of chicks initiated feeding at 25 h post-hatch, increasing to 50% by 31 h post-hatch. However, the crop palpation method used by Boyner et al. (2021) is relatively insensitive as early ingested feed particles are minute and dissolve rapidly, likely leading to an underestimation of the timing of first feed intake. Our results are supported by earlier research, showing that chicks consumed an average of 1.5 g of feed within the first 24 hours (Pinchasov and Noy, 1993). Notably, unchanged ghrelin mRNA expression in the feed-only treatment suggests that feed intake in the absence of water may be minimal. Providing water alone can help reduce BW loss and dehydration, but does not appear to suppress ghrelin mRNA expression. This suggests that water provision alone may be insufficient to provide the prandial or nutrient-related signal required to reduce ghrelin expression, most likely because ghrelin is primarily responsive to changes in feeding status and feed was not available in W. In addition, the elevated plasma CORT concentrations observed in W further support the importance of providing both feed and water during the post-hatch period.

Providing both feed and water immediately after hatching reduces CORT levels during the post-hatch period. Similar reductions in CORT levels following EF have been reported previously (van de Ven et al., 2011; Wijnen et al., 2022). However, other studies did not observe such effects (Gonzales et al., 2003; Tong et al., 2015; van de Ven et al., 2013) and highlight some inconsistency in literature. Delayed access to feed and water, however, induces only temporary increases in CORT, as elevated levels declined rapidly following feed access (Wijnen et al., 2022). Further, no long-term elevations were detected at later ages (Wijnen et al., 2022). Importantly, caution is warranted when interpreting fluctuations in plasma CORT during feed deprivation as a direct indicator of a negative affective state, as corticosterone is also a physiological hormone involved in maintaining homeostasis during periods of nutritional imbalance through regulation of carbohydrate, protein, and lipid metabolism. Behavioural assessments would be required to determine whether changes in plasma CORT post-hatch reflect an altered affective state or are associated with nutritional imbalance. Unfortunately these were not included in the present study.

A limitation of the present study is that only chicks originating from the middle 60% of the hatch window were included. While this improved experimental uniformity and facilitated the practical execution of the study, it may limit the direct relevance of the findings to commercial hatcheries. Further research including the full hatch window is therefore recommended.

Conclusion

Taken together, our findings imply that access to both feed and water is required to stimulate feeding post-hatch. When both resources are available, our physiological and growth data suggest that chicks initiate feeding and drinking between 14 and 24 hours post-hatch. This timing was inferred from indirect physiological indicators of water and feed intake, including decreased plasma osmolality and reduced ghrelin mRNA expression, respectively. This resulted in a gradual body weight gain between 24 and 48 hours post-hatch. Provision of water alone attenuated body weight loss compared with total deprivation or feed-only provision. Moreover, withholding water for more than 48 hours post-hatch resulted in a more pronounced state of dehydration relative to chicks receiving water immediately post-hatch. Absolute yolk sac utilization was not affected by feed and/or water availability. Plasma corticosterone levels were elevated at 36 and 48 hours post-hatch in water-, feed- or totally-deprived chicks, whereas the lowest levels were observed when both feed and water were available. However, these differences may reflect a multifactorial interplay between stress-related and metabolic responses and therefore warrant further investigation.

Funding

This study received the financial support of VLAIO (Flemish Innovation & Entrepreneurship) through the project HBC.2021.1063

Disclosures

The authors declare that they have no known competing financial interests or personal relationships that could have appeared to influence the work reported in this paper.