Animals and nutritional treatment

Animal care and use protocols were approved by the Ethical Animal Committees of Federal Fluminense University (CEUA/UFF 1032/2018) and adhered with the National Institute of Health Guide for the Care and Use of Laboratory Animals.^{Reference Albus10} Female Wistar rats from the Experimental Nutrition Laboratory of Federal Fluminense University (3 months of age) were mated and then individually housed in cages with food (standard chow or cocoa powder-supplemented chow), and tap water was provided ad libitum. The animals were maintained in a temperature- and light-controlled environment (24°C ± 2°C, 12 h light, 12 h dark, lights on at 07:00 am). After birth, litter numbers were adjusted for optimal lactotrophic performance (n = 6 male pups/mother)^{Reference Langley-Evans, Phillips and Benediktsson11}, and at weaning, the animals were randomly separated into four experimental groups (n = 6 males/group) according to perinatal and postweaning feeding: control (C) group – offspring of mothers fed standard diet, weaned at 21 d, and received the same diet in the postnatal life; cocoa control (CCa) group – offspring of mothers fed standard diet supplemented with 10% cocoa powder, weaned at 21 d, and given the same diet in the postnatal life; early weaning (EW) group – offspring of mothers fed standard diet, weaned at 18 d, and given the same diet in the postnatal life; and cocoa early weaning (EWCa) group – offspring of mothers fed standard diet supplemented with 10% cocoa powder, weaned at 18 d, and given the same diet in the postnatal life. The diet supplemented with cocoa powder was handled in the laboratory with the following components: 90 g of standard diet (NUVILAB®, Curitiba, Brazil) and 10 g of cocoa powder (Armazém®, Rio de Janeiro, Brazil), with a total calorie content of 296.4 kcal/100 g. The animals were followed up to 3 months of age. The literature shows that the increase of 10% of cocoa powder in the diets is within the standards used in animals.^{Reference Álvarez-Cilleros, Ramos and López-Oliva12,Reference Álvarez-Cilleros, López-Oliva and Martín13}

Dietary intake, body weight, and blood glucose

Food intake and body mass were determined daily and weekly, respectively. To determine weight gain, the weight of the animals shortly after early weaning was subtracted from the final weight at the age of 90 d. For serum glucose analysis, the animals were maintained at an 8-h fasting period, and blood glucose concentrations were measured using a glucometer (Accu Check, Roche, SP, Brazil). Blood samples were taken from the tail vein of the rats.

Euthanasia and organ collection

At euthanasia, the animals were deprived of food for a period of 8 h and anesthetized with an intraperitoneal injection of xylazine (0.1 mg/kg) and ketamine (0.8 mg/kg). Blood was collected by cardiac puncture (right atrium) and centrifuged at 3500 rpm and 4°C for 15 min to obtain the serum (SIGMA® 4K15 centrifuge, Germany), which was frozen at −80°C for future analysis of total cholesterol (TC) (Monoreagent Cholesterol – K083; Bioclin®), high-density lipoprotein cholesterol (HDL-c) (Direct HDL – K071; Bioclin®), and triacylglycerol (TAG) (Triglycerides Monoreagent – K117; Bioclin®). For the determination of the low-density lipoprotein cholesterol (LDL-c) fraction, Friedewald equation (LDL = TC – HDL – (TAG/5) was used.^{Reference Friedewald, Levy and Fredrickson14}

Subsequently, the testicles were dissected, weighed on an analytical balance (Sartorius TE214S, Chicago, IL, USA), and fixed in Bouin for 24 h. The male gonad was then cleaved and fixed again (4% buffered formalin in phosphate buffer of pH 7.2) for a period of 48 h. In addition, the retroperitoneal, epididymal, and brown fats were excised, weighed separately with the same scale, and fixed in buffered formalin.

Body composition evaluation

After anesthesia, the animals underwent dual-energy X-ray absorptiometry (DXA) using the GE Lunar DXA 200, 368 instrument (Lunar, WI, USA) with specific software (Bis 2008, version 12.20 GE Healthcare, Milwaukee, WI, USA). Total body mass (g), lean mass (g), fat mass (g), body fat percentage (%), lean body mass (g), trunk fat mass (g), body fat percentage trunk (%), and bone analysis [bone mineral density (g/cm²), bone mineral content (g), and bone area (cm²)] were measured for each animal. The assessment was blind, as the DXA technician was unaware of the experimental protocol.

Testicular morphometry

The seminiferous tubule diameter and the seminiferous epithelium height were determined by morphometry using the Image J® Software (Image Processing and Analysis in Java, Bethesda, MD, USA). Briefly, 25 microscopic fields were analyzed per animal, making up 125 seminiferous tubules, according to techniques previously described.^{Reference Ribeiro, De Souza and Medeiros15,Reference De Souza, Silva and Cortez16} Surface density (Sv; proper tunic, seminiferous epithelium, tubular lumen, and stroma) was estimated by point count: Sv = Pp/Pt, where Pp is the number of points that touch the structure and Pt is the number of total points of the test system.^{Reference De Souza, Silva and Cortez16} Sertoli and Leydig cells were counted directly through J® Software using the cell counter tool, at 600× magnification.^{Reference Shekarforoush, Ebrahimi and Hoseini17–Reference Kotsampasi, Balaskas, Papadomichelakis and Chadio19}

Statistical analysis

All data were tested for normal distribution curve and homogeneity of variance using the Kolmogorov–Smirnov test, with one-way analysis of variance and Bonferroni post hoc test. For parameters that did not reach normal distribution, the Kruskal–Wallis test with Dunn post hoc test was used (GraphPad Prism version 5.03 for Windows; GraphPad Software, San Diego, CA, USA). Results are presented as mean ± standard deviation (SD). P ≤ 0.05 was considered statistically significant.