Supplementing entire male pig diet with hydrolysable tannins: Effect on carcass traits, meat quality and oxidative stability

Meat Science 133 (2017) 95–102

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Supplementing entire male pig diet with hydrolysable tannins: Effect on carcass traits, meat quality and oxidative stability

MARK

Vida Rezara, Janez Salobira, Alenka Levarta, Urška Tomažinb, Martin Škrlepb, Nina Batorek Lukačb, Marjeta Čandek-Potokarb,c,⁎ a b c

Department of Animal Science, Biotechnical Faculty, University of Ljubljana, Groblje 3, 1230 Domžale, Slovenia Agricultural Institute of Slovenia, Hacquetova Ulica 17, 1000 Ljubljana, Slovenia University of Maribor, Faculty of Agriculture and Life Sciences, Pivola 10, 2311 Hoče, Slovenia

A R T I C L E I N F O

A B S T R A C T

Keywords: Pig Tannin Carcass Meat Oxidative stability

The purpose of the present study was to investigate the potential impact on carcass and meat quality of a sweet chestnut wood extract (SCWE)diet supplement for pigs, in particular on oxidative stability and fatty acid composition. Entire (non-castrated) male pigs (n = 24) were assigned to treatment groups within litter and offered one of 4 finisher diets on an ad libitum basis: T0 (control), T1, T2 or T3, supplemented with 0, 1, 2 or 3% of commercially available SCWE, respectively. The highest SCWE supplementation reduced carcass fat deposition and water holding capacity of meat (higher thawing loss). In fresh meat, SCWE supplementation increased lipid (malondialdehyde) and protein oxidation (carbonyl groups in myofibril isolates). With regard to fat tissue, SCWE supplementation increased the proportion of polyunsaturated fatty acids.

1. Introduction Tannins are secondary plant metabolites with great structural diversity (classified as hydrolysable, condensed or complex tannins) exerting different physiological effects according to their form, the amount ingested and the animal species involved (for detailed review see Mueller-Harvey, 2006). In certain amounts tannins are considered anti-nutritive substances because they precipitate proteins, inhibit digestive enzymes and affect the utilisation of vitamins and minerals (Chung, Wei, & Johnson, 1998). In addition they reduce feed palatability and consequently feed intake and animal performance (Jansman, 1993; Mueller-Harvey, 2006). On the other hand, certain tannins are, due to their antimicrobial properties, commonly fed to monogastric animals as antihelmintic, antimicrobial and antiviral agents, and as supportive treatment for diarrhoea (Mueller-Harvey, 2006; Redondo, Chacana, Dominguez, & Fernandez Miyakawa, 2014). Tannins may work as antioxidants to scavenge free radicals (Riedl, Carando, Alessio, McCarthy, & Hagerman, 2002). Sweet chestnut (Castanea sativa Mill.) wood extract (SCWE), consisting mainly of hydrolysable tannins, was shown to possess reducing and antioxidant capacity in in vitro trials (Lampire et al., 1998), was able to reduce in vivo oxidative stress in pigs and poultry (Frankič & Salobir, 2011;Voljč, Levart, Žgur, & Salobir, 2013) and increased oxidative stability of meat through preserving vitamin E (Voljč et al., 2013). In pigs, it was

⁎

demonstrated that hydrolysable tannins, besides reducing total protein digestibility (Antongiovanni, Minieri, & Petacchi, 2007; Salobir, Kostanjevec, Štruklec, & Salobir, 2005), inhibit protein fermentation in the colon (Biagi, Cipollini, Paulicks, & Roth, 2010) and reduce intestinal production of skatole, a boar taint compound (Čandek-Potokar et al., 2015). Because of the possible effects on the digestion of protein and other nutrients, and due to its antioxidant properties, tannin supplementation is likely to affect carcass and meat quality. However, scientific literature on this topic is still lacking, especially in relation to pigs. Apart from the research dealing with pigs fed with acorns and chestnuts (García-Valverde, Nieto, Lachica, & Aguilera, 2007; Pugliese et al., 2009; Tejerina, García-Torres, Cabeza de Vaca, Vázquez, & Cava, 2011), other literature data refers either to ruminants (Luciano et al., 2009, 2011; Vasta, Nudda, Cannas, Lanza, & Priolo, 2008), rabbits (Gai et al., 2009; Liu, Dong, Tong, & Zhang, 2011; Liu, Zhou, Tong, & Vaddella, 2012; Liu et al., 2009) or poultry (Schiavone et al., 2008). Moreover, so far the reported dosages have been low, and despite the fact that pigs (e.g. Iberian) can ingest relatively high levels of tannins, information about the effect of tannins (type and concentration) is lacking. Therefore, the present study was conducted to investigate the potential impact on carcass and meat quality of supplementing the diet of pigs with SCWE rich in hydrolysable tannins, considering also fatty acid composition and oxidative stability.

Corresponding author at: Agricultural Institute of Slovenia, Hacquetova Ulica 17, 1000 Ljubljana, Slovenia. E-mail address: [email protected] (M. Čandek-Potokar).

http://dx.doi.org/10.1016/j.meatsci.2017.06.012 Received 10 April 2017; Received in revised form 8 June 2017; Accepted 22 June 2017 Available online 23 June 2017 0309-1740/ © 2017 Elsevier Ltd. All rights reserved.

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(AOAC method 920.39), crude fibre (fritted glass crucible method, AOAC method 978.10) and crude ash (AOAC method 942.05). Animals were fed experimental diets for 70 days, including a 5-day transitional period. At the age of 193 days and weight of 122.5 kg, the experimental pigs were slaughtered in one batch using routine abattoir procedure (CO2 stunning, dehairing). Feed was withdrawn one day prior to slaughter.

Table 1 Chemical and fatty acid composition (% of total fatty acids) of feed mixtures. Treatment groupa

Dry matter (g/kg) Crude ash (g/kg) Crude protein (g/kg) Crude fat (g/kg) Crude fibre (g/kg) Nitrogen free extract (g/kg) α-tocopherol (mg/kg) γ-tocopherol (mg/kg) Main fatty acid composition (% of the total fatty acids)b C 12:0 C 14:0 C 16:0 C 18:0 Σ C18:1 C 18:2 n − 6 C 18:3 n − 3 Σ SFA Σ MUFA Σ PUFA Σ n − 3 PUFA Σ n − 6 PUFA Σ n − 6/Σ n − 3 PUFA

T0

T1

T2

T3

892 48 175 26 52 593 67.3 20.9

885 45 169 29 47 595 70.4 21.3

881 46 166 28 49 591 71.4 21.7

885 45 164 27 50 599 69.6 21.9

< 0.01 0.09 14.0 2.2 27.9 51.2 2.7 17.4 28.7 53.9 2.7 51.3 19.2

< 0.01 0.08 13.7 2.4 27.7 50.9 3.3 17.2 28.5 54.2 3.3 50.9 15.5

< 0.01 0.08 13.6 2.3 28.0 51.1 3.1 17.1 28.7 54.2 3.1 51.1 16.4

< 0.01 0.08 13.7 2.3 27.9 51.0 3.1 17.2 28.6 54.1 3.1 51.0 16.6

2.2. Carcass and meat quality measurements After the slaughter, pigs were eviscerated, leaf (i.e. subperitoneal) fat was removed, carcasses split apart, weighed and classified by the official classification body, using a method approved for Slovenia (OJ EU L56/28, 2008). The method consists of measuring minimal fat thickness over gluteus medius muscle (backfat thickness) and the shortest distance between cranial end of gluteus medius and dorsal edge of vertebral canal at the carcass split line with a digital caliper and enables calculation of lean meat content. Measurement of pH (pH 3) was taken in longissimus lumborum muscle (LL) at the level of last rib 3 h post mortem using a MP120 Mettler-Toledo pH meter (Mettler-Toledo GmbH, Schwarzenbach, Switzerland). The carcasses were cooled overnight at 0–2 °C until the internal temperature dropped below 7 °C. A day after the slaughter the carcasses were cut at the level of last rib perpendicularly to the spine. The measurements of CIE L*, a*, b* colour parameters (using Minolta Chroma Meter CR-300, Minolta Co. Ltd., Osaka, Japan) and ultimate pH were performed on a freshly cut surface of LL. A digital photo of the cross-section was taken and the measurements of loin eye area and area of the corresponding fat performed using LUCIA.NET 1.16.5 software (Laboratory Images s.r.o., Prague, Czech Republic) performed as described in Batorek et al. (2012). Caudally from last rib, two 2.5 cm thick chops of LL were taken, trimmed of epimysium and external fat and used for the determination of drip loss, chemical composition, thawing loss, cooking loss and shear force. Drip loss was determined according to the EZ method (Christensen, 2003). In short, two cylindrical samples of LL were excised, weighed and stored in plastic containers at 4 °C. Drip loss was expressed as the difference (%) from the initial sample weight after 24 and 48 h. For determination of chemical composition, LL samples were minced and the moisture, intramuscular fat (IMF) and protein content were determined using near-infrared spectral analysis (NIR Systems 6500, Foss NIR System, Silver Spring, MD, USA) using internal calibration (Prevolnik et al., 2005). The second LL chop was weighed, vacuum packed and frozen at − 20 °C until analysis. Samples were thawed (overnight at 4 °C), gently drained with a paper towel, weighed and the difference in weights used for thawing loss calculation. The same samples were then cooked in a thermostatic bath (ONE 7-45, Memmert GmbH, Schwabach, Germany) until the internal temperature reached 72 °C, drained and reweighed for cooking loss evaluation, and cooled overnight at 4 °C. The next day, shear force was measured on two 2.5-cm-wide cylindrical cores excised from the sample using TA Plus texture analyzer (Ametek Lloyd Instruments Ltd., Bognor Regis, UK) equipped with a 60° V-shaped blade at a crossheaf speed of 3.3 mm/s.

a The control group (T0) received feed without supplementation, while the experimental groups T1, T2 and T3 were offered the same diet supplemented with 1%, 2% and 3% of commercially available sweet chestnut wood extract, respectively. Diet T0 was composed of maize (62%), soya meat (13%), wheat meal (8%), rapeseed meal (7%), sunflower meal (5%), molasses (2%), CaCO3 (1.1%), NaCl (0.6%), lysine (1%), methionine (0.3%), Ca(H2PO4)2 (0.17%). b Only predominant fatty acids are listed, but the sum of saturated fatty acids (SFAs), monounsaturated fatty acids (MUFAs) and polyunsaturated fatty acids (PUFAs) are computed using all analysed fatty acids.

2. Materials and methods 2.1. Animals and diets Experimental design, animals and feed composition are described in detail elsewhere (Čandek-Potokar et al., 2015). As explained therein, the work was undertaken with full owner compliance and within the normal running of the farm. The study was performed following the Slovenian law on animal protection (Zakon o zaščiti živali, 2007) and was not subject to ethical protocols according to Directive 2010/63/EU (2010), i.e. approved feed additives were used (European Union Register of Feed Additives, 2013) and tissue samples were taken after the slaughter. Briefly, 24 crossbred (Large White × Landrace) entire male pigs were allocated within litters to four treatment groups, housed individually with ad libitum access to feed and water. All the animals were fed a commercial feed mixture (Table 1) that was calculated to contain 13.2 MJ ME/kg and to meet requirements according to NRC (2012). The control group (T0) received feed without supplementation, while the experimental groups T1, T2 and T3 were offered the same feed supplemented with 1%, 2% and 3% of commercially available SCWE (Farmatan®, Tanin Sevnica d.d., Sevnica, Slovenia), respectively. Sweet chestnut wood extract is rich in hydrolysable tannins, mainly gallotannins (Bee et al., 2016; Biagi et al., 2010). The analysis of SCWE for total phenols – determined by the Folin-Ciocalteau colourimetric method (Rigo et al., 2000) and expressed as gallic acid equivalents – showed the content of 43.6%. Feed samples were taken at the beginning and at the end of the experiment, pooled and used for proximate analysis and determination of fatty acid composition. Proximate analysis (moisture, crude ash, crude protein, crude fat, crude fibre) of feed was performed according to standard procedures (AOAC, 2000): dry matter (in oven drying at 95–100 °C AOAC method 934.01), crude protein (the copper catalyst Kjeldahl method AOAC method 984.13), crude fat

2.3. Preparation of meat and subcutaneous fat samples for chemical analysis Muscle and fat samples were trimmed of other tissues. Each muscle sample was divided into two parts: one was homogenised in liquid nitrogen, the other was transferred to 50-mL polypropylene containers with covers, cooked in a water bath (LCS 0081 Lauda, Bartelt GmbH, Graz, Austria) for 60 min at 90 °C, and then cooled to room temperature in a cold water bath. All fat and muscle samples were frozen in liquid nitrogen and homogenised using a laboratory knife mill (Grindomix GM200, Retsch GmbH and Co., Haan, Germany). Concentration of carbonyls was determined in myofibril isolates from fresh muscle 96

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50% trichloroacetic acid. After centrifugation (4000 × g for 15 min at 4 °C), the pellets were washed three times with a 1 ml of mixture of ethanol:ethyl acetate (1:1) to eliminate traces of residual DNPH. The pellets were afterwards dissolved in 2 ml of 6 M guanidine HCl with 20 mM sodium phosphate buffer (pH 6.5) and centrifuged for 15 min at 4000 ×g. The protein and carbonyl concentrations were spectrophotometrically determined in the supernatant (BioSpectrometer Fluorescence, Eppendorf, Hamburg, Germany). The concentration of protein was calculated from absorbance at 280 nm according to the standard concentrations of BSA in 6 M guanidine HCl while the concentration of carbonyls was measured in the samples treated with DNPH at 370 nm (considering that the extinction coefficient for DNPH at 370 nm is 21/mM− 1 cm− 1).

samples, while tocopherols, malondialdehyde (MDA), and antioxidant capacity of the lipid-soluble compounds (ACL) were determined in both fresh and thermally treated meat. Fatty acid composition of meat (fresh) and subcutaneous fat were also determined. 2.3.1. Determination of vitamin E (α- and γ-tocopherol) Concentrations of tocopherols in feed and meat samples were measured according to the methodology of Abidi and Mounts (1997) and Rupérez, Martín, Herrera, and Barbas (2001). Samples were treated with ethanol and the tocopherols were extracted using hexane. Hexane phases were transferred to clean tubes and evaporated under a gentle stream of nitrogen. Residues were subsequently dissolved in ethanol. Samples were transferred to autosampler vials and analysed by reversephase HPLC using a Prodigy 5 μ ODS(2) column (250 × 4.6 mm, Phenomenex Inc., USA). The mobile phase consisted of methanol and had a flow rate of 1.5 ml/min. Results of the analysis were evaluated using the Agilent HPLC (Agilent Technologies, Wilmington, DE, USA). The instrument was calibrated using external standards (Tocopherol set, Sigma-Aldrich Co.).

2.3.5. Fatty acids composition The fatty acid composition of the diets, intramuscular fat and subcutaneous fat was determined using gas chromatography following transesterification of lipids (Fidler, Salobir, & Stibilj, 2000). Approximately 0.5 g of each sample was transmethylated in situ (Park & Goins, 1994) using 0.5 M NaOH in methanol followed by 14% BF3 in methanol. Fatty acid methyl ester (FAME) was extracted using hexane. For FAME separation, an Agilent GC (Agilent, Santa Clara, CA; USA) equipped with an Omegawax 320 column (30 m × 0.32 mm i.d. × 0.25 μm; Supelco, Bellefonte, PA, USA) and FID detector was used. An Agilent GC ChemStation (Agilent, Santa Clara, CA; USA) was used for data acquisition and processing. Individual FAMEs were identified using standard mixtures (Nu Chek Prep Inc., Elysian, MN, USA).

2.3.2. Determination of antioxidant capacity of the lipid-soluble compounds (ACL) The ACL in meat samples was measured using a photochemiluminescence method by PhotoChem® (Analytik Jena, Jena, Germany) and is presented as Trolox equivalent. Lipid- soluble antioxidants from samples were extracted with hexane (0.6 g of homogenised meat sample in 500 μl of water and 500 μl of hexane), mixed on a vortex and centrifuged (10.000 ×g, 10 min, 4 °C). The supernatants were analysed in accordance with ACL Kit protocol (Analytik Jena, Jena, Germany). Briefly, 10 μl of supernatant was mixed with 2300 μl of R1 (methanol), 200 μl of R2 (buffer) and 25 μl or R3 (photosensitizer and detection reagent containing luminol), and analysed. Inhibition of luminescence signal (integration over 250 s) in comparison to blank was calculated (PCLsoft). For calibration of instrument, trolox was used in 0.5 to 2.0 nmol range.

2.4. Statistical analysis An analysis of variance (one-way ANOVA) for the effect of treatment group was performed by MIXED procedure of SAS statistical software 9.2 (2011) (SAS Inst., Inc., Cary, NC, USA). In the case of significant treatment group effect, the differences (between leastsquares means) were tested with the Dunnett test (i.e. comparing each treatment group with the control) and in the case of significance, results are presented in Figs. 1–4. Additionally, carcass weight and backfat thickness -were tested as a covariate in the case of carcass traits and fatty acid composition, respectively; results are not shown, but are used for discussion.

2.3.3. Determination of malondialdehyde (MDA) Oxidation of meat lipids was monitored by measurement of MDA concentration in individual fresh and heat-processed meat samples. A modified version of the methodology of Vila, Jaradat, Marquardt, and Frohlich (2002) was implemented to determine MDA concentration in samples using HPLC. Homogenised samples (0.3 g) were mixed with 1.5 ml of 2.5% trichloroacetic acid in 2 ml microcentrifuge tubes, left for 10 min and then centrifuged (15,000 × g for 15 min at 4 °C). One ml of supernatant was mixed with 1.5 ml of 0.6% thiobarbituric acid and heated at 90 °C for 1 h. After cooling, the samples were filtered through Millipore filters (0.22 μm) into auto-sampler vials. The Agilent HPLC system was used with a 1260 Infinity fluorescence detector. For separation, a reverse-phase HPLC chromatography column (HyperClone 5 u ODS (CI8) 120A, 150 × 4.6 mm; Phenomenex Inc., USA) and CI8 ODS guard column (4 × 3 mm; Phenomenex Inc., USA) were used. The mobile phase consisted of a 65% 50 mM KH2P04 buffer (pH 6.9) and 35% methanol. The mobile phase flow rate was 1.0 ml/min.

3. Results and discussion 3.1. Carcass traits There was no major effect of the treatment groups noted for carcass traits (Table 2) except for the reduced fat deposition in the group with the highest (3%) addition of tannin extract (Fig. 1). In T3 group, backfat Table 2 Carcass traits - least squares means and effect of treatment group. Treatment groupa

2.3.4. Concentration of carbonyl groups in myofibril isolates Protein oxidation was measured according to the method of Oliver, Ahn, Moerman, Goldstein, and Stadtman (1987) as modified by Mercier, Gatellier, Viau, Remignon, and Renerre (1998) in myofibril isolates which were prepared according to Pietrzak, Greaser, and Sosnicki (1997). The concentration of carbonyl groups (nmol/mg of protein) was detected by 2,4-dinitrophenylhydrazine (DNPH) to form protein hydrazones. Two aliquots (300 μl) of myofibrillar suspension were treated with 1 ml of 2 N HCl (to determine the concentration of proteins), while two aliquots were treated with an equal volume of 0.2% (w/v) DNPH in 2 N HCl. Samples were incubated for 1 h at room temperature under agitation and afterwards precipitated by 300 μl of

Warm carcass weight, kg Dressing percentage, % Loin thickness, mm Backfat thickness, mm Lean meat content, % Loin eye area, cm2 LL overlying fat area, cm2

T0

T1

T2

T3

SEM

P-value

93.6 75.2 74.7 11.2 61.9 47.8 13.4

95.4 76.5 75.7 11.0 62.5 50.0 13.5

96.7 76.8 73.8 10.6 62.1 51.1 13.2

86.3 74.8 72.5 7.5 64.2 47.8 10.8

3.40 0.62 2.34 0.77 0.74 1.99 0.54

0.18 0.11 0.81 0.001 0.15 0.64 0.005

LL – longissimus lumborum muscle, SEM – standard error of mean, n = 24. a The control group (T0) received feed without supplementation, while the experimental groups T1, T2 and T3 were offered the same diet supplemented with 1%, 2% and 3% of commercially available sweet chestnut wood extract, respectively.

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Fig. 1. Effect of tannin supplementation on carcass and meat quality traits. Only traits significantly affected (P < 0.05) by dietary treatment are depicted. The values shown represent differences between the control and supplemented groups. The levels of significance are denoted as: NS – P > 0.10; †P > 0.05; *P < 0.05; **P < 0.01. T0 – control group, no sweet chestnut wood extract supplement; T1–1% sweet chestnut wood extract; T2–2% sweet chestnut wood extract; T3–3% sweet chestnut wood extract.

thickness and fat area over LL were reduced by 32% and 20%, respectively (P < 0.05) compared to T0 group. The results reflect the differences in fattening performance as previously reported (ČandekPotokar et al., 2015). However, it is worth noting that although smaller, the difference in fat deposition remained significant also when groups were compared at the same carcass weight, whereas no effect was noted for muscular deposition. Previous findings suggest that tannins not only affect food consumption and digestion but also decrease the efficiency of converting the absorbed nutrients to new body substances (Chung, Wong, Wei, Huang, & Lin, 1998). Since the concentration of tannins in our experiment was two to six times higher than in the previously mentioned studies it was interesting to observe rather limited impact on growth performance and body composition. It is possible that negative effects on nutrient utilisation were counterbalanced by the positive influence on gut health (Biagi et al., 2010; Chung, Wei, & Johnson, 1998) or by adaptation ability of salivary glands (Čandek-Potokar et al., 2015) producing increased amounts of tannin-binding proline-rich proteins (Cappai, Wolf, Pinna, & Kamphues, 2013). Recent publications, testing similar supplementations with hydrolysable tannin extract, also show no major effect on pig performance (Bee et al., 2016; Bilić-Šobot et al., 2016), proving that pigs are relatively resistant to high doses of ingested tannins. As for the effect of dietary tannin addition on body fatness (i.e. reducing fat deposition), the two mentioned studies showed no effects. However, numerous studies on various dietary polyphenols show their anti-obesity potential (Lei et al., 2007), interfering with biosynthesis of cholesterol and utilisation of lipids (Axling et al., 2012) and affecting carbohydrate metabolism (Hanhineva et al., 2010). As reviewed by Wang et al. (2014), in vitro cellular studies demonstrated that polyphenols reduce adipocyte viability and preadipocyte proliferation, suppress adipocyte differentiation and triglyceride accumulation, and stimulate lipolysis and β-oxidation. In compliance, studies on laboratory animals also showed lower body weight, fat mass, and triglyceride concentration through enhancing energy expenditure and fat utilisation in relation to consuming polyphenols.

Table 3 Meat quality traits of longissimus lumborum muscle - least squares means and effect of treatment group. Treatment groupa

Water, % Intramuscular fat, % Protein, % pH 3 h pH 24 h Drip loss 24 h, % Drip loss 48 h, % Thawing loss, % Cooking loss, % Shear force, N L* (lightness) a* (redness) b* (yellowness)

T0

T1

T2

T3

SEM

P-value

74.1 1.23 24.1 6.04 5.29 5.2 7.2 11.0 35.4 185.3 57.9 7.6 4.8

74.9 1.07 23.8 5.87 5.34 5.7 8.2 14.7 35.4 196.1 55.6 7.1 3.9

74.1 1.07 23.9 5.71 5.33 6.2 8.8 15.4 34.0 161.6 56.1 7.9 4.3

74.3 1.06 23.7 5.89 5.30 6.5 8.9 15.7 33.5 170.5 57.5 8.0 4.9

0.3 0.09 0.13 0.11 0.03 0.69 0.81 1.17 0.89 13.0 0.8 0.5 0.4

0.47 0.49 0.14 0.31 0.48 0.64 0.47 0.044 0.36 0.30 0.20 0.56 0.33

a The control group (T0) received feed without supplementation, while the experimental groups T1, T2 and T3 were offered the same diet supplemented with 1%, 2% and 3% of commercially available sweet chestnut wood extract, respectively. SEM – standard error of mean, n = 24.

Table 4 Concentration of α- and γ-tocopherol, antioxidant capacity of lipid-soluble compounds (ACL), malondialdehyde (MDA) and carbonyls in meat (longissimus lumborum muscle) least squares means and effect of treatment group. Treatment groupa

α-Tocopherol, μg/100 g muscle Fresh Cooked γ-Tocopherol, μg/100 g muscle Fresh Cooked ACL, nmol/g muscle Fresh Cooked MDA, nmol/g muscle Fresh Cooked Carbonyls, nmol/mg proteins

3.2. Meat quality Dietary treatments with tannins did not result in any major differences in observed physical-chemical properties of meat quality or chemical composition (Table 3). Although carcass adiposity of T3 group was significantly lower than in other groups, no effect on IMF content was observed. This result could be related to the fact that IMF was generally very low, as experimental pigs were entire (non-castrated) males which exhibit higher leanness than their castrated counterparts (Batorek et al., 2012). The only statistically important difference observed among the groups was in thawing loss (Fig. 1); in T3 group it was 43% higher than in T0 group (P < 0.05). The thawing loss in T1 and T2 was also notably higher than in T0 (34 and 40% respectively), and close to significance (P = 0.16 and P = 0.09, respectively). Although the differences were less notable (P > 0.10), other water holding capacity (WHC) indicators (drip loss after 24 and 48 h) showed the same trend, (i.e. values increasing with higher tannin doses).

T0

T1

T2

T3

SEM

P-value

268.2 242.2

267.9 256.2

241.1 258.2

202.5 252.9

21.3 16.3

0.036 0.91

8.19 7.58

9.15 9.45

7.60 8.38

6.31 7.63

0.69 0.81

0.058 0.36

3.69 3.27

3.73 3.11

3.77 3.06

3.87 3.12

1.10 0.24

0.61 0.94

0.23 7.38 1.20

0.28 5.92 1.83

0.26 3.33 1.95

0.39 5.14 2.58

0.03 0.60 0.13

0.014 0.002 < 0.001

SEM – standard error of mean, n = 24. a The control group (T0) received feed without supplementation, while the experimental groups T1, T2 and T3 were offered the same diet supplemented with 1%, 2% and 3% of commercially available sweet chestnut wood extract, respectively.

3.3. Tocopherols, ACL, MDA and carbonyl levels in meat The effect of dietary tannin supplementation was observed on αand γ-tocopherol, MDA and carbonyl concentrations of LL (Table 4). However, it should be emphasised that the effect of treatment on α- and γ-tocopherol concentrations was noted only in T3 and is presumably an artefact, as it was not confirmed in cooked samples (higher values than 98

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Fig. 2. Effect of tannin supplementation on indicators of muscle oxidation. Only traits significantly affected (P < 0.05) by dietary treatment are depicted. The values shown represent differences between the control and supplemented groups. The levels of significance are denoted as: NS – P > 0.10; *P < 0.05; **P < 0.01. T0 – control group, no sweet chestnut wood extract supplement; T1–1% sweet chestnut wood extract; T2–2% sweet chestnut wood extract; T3–3% sweet chestnut wood extract; MDA – malondialdehyde.

threshold level (0.50 mg/kg) for rancidity detection by trained sensory panellists (Dunshea, D'Souza, Pethick, Harper, & Warner, 2005). Low levels of MDA in fresh meat samples can be related to low amounts of polyunsaturated fatty acids (PUFAs) and sufficient vitamin E concentration for adequate PUFA protection (the minimal physiological requirement of vitamin E needed for protection of PUFA is 112–131 μg/ 100 g; Muggli, 1994). Increased MDA concentration along with increased concentration of carbonyl groups indicates prooxidative effect of tannin supplementation denoting that meat oxidation could be further increased during storage (Filgueras et al., 2010; Mercier et al., 1998) or processing (i.e. in dry-cured meat products) (Koutina, Jongberg, & Skibsted, 2012). As suggested by Traore et al. (2012) and Lonergan, Huff-Lonergan, Rowe, Kuhlers, and Jungst (2001), inferior WHC may be directly associated with higher oxidation level of either proteins (i.e. carbonyls) or lipids (i.e. MDA), leading to protein denaturation, thus lowering their ability to bind water (Joo, Kauffman, Kim, & Park, 1999; Offer & Knight, 1988), as can be observed also in the present study. With regard to the possible effects of tannins on vitamin E content and oxidative stability of meat in monogastrics, most research was done in chickens and rabbits and shows mostly no effect, or some positive effects, on vitamin E concentration and oxidative stability of meat. However, the concentrations of tannins used were much lower in comparison to our experiment. Liu et al. (2009) found significantly lower iron-induced lipid oxidation in rabbit meat with 0.5% dietary tannin supplementation. In contrast, Schiavone et al. (2008) observed no effect of dietary tannin supplementation (0.15 to 0.25%) on lipid oxidation in raw or heat-treated breast muscle of broilers. Similarly, Voljč et al. (2013) showed that tannins at 0.3% in broiler diet had no effect on concentration of vitamin E in raw, and MDA in raw and cooked, breast muscle. According to Surai (2014), polyphenols do not seem to offer in vivo antioxidant protection similar to that of vitamin E, since they are not efficiently absorbed from the gastrointestinal tract, they are quickly metabolised and excreted from the organism. However, Luciano et al. (2009, 2011) showed significant accumulation of phenolic compounds after dietary supplementation of condensed tannin-rich quebracho in lambs, resulting in improved overall muscle antioxidant status but with no effect on lipid oxidation.

Table 5 Fatty acid composition (% of total fatty acids) of subcutaneous and intramuscular fat least squares means and effect of treatment group. Treatment groupa T0

T1

T2

T3

SEM

P-value

Subcutaneous fat Σ SFA Σ MUFA Σ PUFA Σ n − 6 PUFA Σ n − 3 PUFA n − 6/n − 3 PUFA Σ LC PUFA Σ LC n − 6 PUFA Σ LC n − 3 PUFA Σ LC n − 6/Σ n − 3 PUFA

36.69 44.01 19.30 18.27 0.97 18.86 1.48 1.25 0.23 5.45

38.14 41.05 20.82 19.47 1.29 15.10 1.49 1.22 0.27 4.53

37.90 41.28 20.82 19.53 1.24 15.81 1.49 1.23 0.26 4.73

33.90 41.30 24.81 23.33 1.41 16.60 1.81 1.50 0.31 4.90

0.75 0.58 0.86 0.80 0.06 0.21 0.07 0.06 0.01 0.08

0.003 0.005 0.002 0.002 < 0.001 < 0.001 0.006 0.006 0.003 < 0.001

Intramuscular fat Σ SFA Σ MUFA Σ PUFA Σ n − 6 PUFA Σ n − 3 PUFA Σ n − 6/Σ n − 3 PUFA Σ LC PUFA Σ LC n − 6 PUFA Σ LC n − 3 PUFA Σ LC n − 6/Σ n − 3 PUFA

36.81 41.24 21.94 20.90 1.03 20.26 6.58 5.89 0.69 8.51

38.09 37.99 23.91 22.68 1.23 18.36 7.09 6.29 0.80 7.85

38.54 38.53 22.92 21.79 1.11 19.49 7.03 6.28 0.75 8.35

37.16 36.81 26.03 24.71 1.31 18.84 7.87 7.00 0.88 7.96

0.40 1.36 1.55 1.50 0.07 0.47 0.66 0.60 0.06 0.25

0.029 0.17 0.32 0.35 0.039 0.056 0.59 0.63 0.24 0.23

a The control group (T0) received feed without supplementation, while the experimental groups T1, T2 and T3 were offered the same diet supplemented with 1%, 2% and 3% of commercially available sweet chestnut wood extract, respectively. SEM - standard error of mean, SFA – saturated fatty acids, MUFA – monounsaturated fatty acids, PUFA – polyunsaturated fatty acids, LC – long chain (C ≥ 20), n = 24.

in fresh meat). The muscle antioxidant capacity indicator (ACL) was not affected by treatment, whereas increased carbonyl groups' concentration and MDA (T3 group only) in fresh meat are indicative of the prooxidant effect of tannin supplementation (Fig. 2). On the other hand, in cooked meat samples there was a significant decrease of MDA in T2 and T3 (55% and 30%, respectively)compared to the control. It should be noted that the observed MDA concentrations detected in fresh meat (from 16.6 μg/kg in T0 to 28.1 μg/kg in T3) and cooked meat (from 0.24 mg/kg in T2 to 0.53 mg/kg in T0) were mainly well below the

3.4. Fatty acid composition of meat and subcutaneous fat tissue Important differences in the fatty acid profile occurred to a much greater extent in subcutaneous fat than in IMF (Table 5). In the case of Fig. 3. Effect of tannin supplementation on fatty acid composition of intramuscular fat (% of the total fatty acids). Only traits, significantly affected (P < 0.05) by dietary treatment are depicted. The values shown represent differences between the control and supplemented groups. The levels of significance are denoted as: NS – P > 0.10; *P < 0.05. T0 – control group, no sweet chestnut wood extract supplement; T1–1% sweet chestnut wood extract; T2–2% sweet chestnut wood extract; T3–3% sweet chestnut wood extract; SFA – saturated fatty acids; PUFA – polyunsaturated fatty acids.

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Fig. 4. Effect of tannin supplementation on fatty acid composition of subcutaneous fat (% of the total fatty acids). Only traits significantly affected (P < 0.05) by dietary treatment are depicted. The values shown represent differences between the control and supplemented groups. The levels of significance are denoted as: NS – P > 0.10; *P < 0.05; **P < 0.01; ***P < 0.0001. T0 – control group, no sweet chestnut wood extract supplement; T1–1% sweet chestnut wood extract; T2–2% sweet chestnut wood extract T3–3% sweet chestnut wood extract; SFA – saturated fatty acids; MUFA – monounsaturated fatty acids; PUFA – polyunsaturated fatty acids; LC – long chain (C ≥ 20).

IMF, dietary treatment influenced total n − 3 PUFAs (mostly marked by α-linolenic acid) that were present in a higher proportion in T3 compared to T0 (Fig. 3). At the same time, an increased proportion of saturated fatty acids (SFAs) was observed in T2. In subcutaneous fat, the effect of hydrolysable tannins supplementation was most pronounced in T3 (Fig. 4). In comparison to T0, the proportion of SFA and monounsaturated fatty acids (MUFAs), especially palmitic (C:16:0) and oleic acid (C18:1 n − 9) was lower in T3, whereas the proportion of almost all fatty acids from n − 6 and n − 3 families was higher (for more details see Supplementary Table 2). i.e. linoleic (C18:2 n − 6), αlinolenic acid (C18:3 n − 3), arachidonic (C20:4 n − 6), docosapentaenoic (C22:5 n − 3) and docosahexaenoic (C22:6 n − 3) acids, and total n − 6 and n − 3 PUFA and long chain (LC) PUFA. In addition, n − 6/n − 3 PUFA and LC-PUFA ratios were also reduced. The effects in T1 and T2 groups followed the same, but less pronounced, trend. In these two groups, a significantly reduced proportion of oleic acid and total MUFA, and increased proportion of α-linolenic acid and total n − 3 PUFA were observed in addition to reduced n − 6/n − 3 PUFA ratio (P < 0.05) (Table 5). It is difficult to know to what extent the tannins could have a direct or indirect effect on the observed fatty acid composition. As the body is able to synthesise only SFAs and MUFAs, the reduced proportion of both in T3 can be simply a consequence of lower feed intake and thus reduced energy availability for fat synthesis. Alternatively, it could be due to lower synthesis of SFA

and MUFA when fat deposition is reduced (Wood et al., 2008), which can also explain increased proportions of linoleic acid in tannin supplemented groups. Similarly in IMF, the gradual increase in linoleic acid proportion can be attributed to low IMF content and consequently more pronounced contribution of PUFA (linoleic acid) rich phospholipid membrane lipids to overall fatty acid composition of IMF. However, when backfat thickness was included in the statistical model, the proportions of SFA and PUFA in subcutaneous fat and n − 3 PUFA proportion in IMF were still affected by the diet, indicating an effect of tannin supplementation regardless of the effect of fat deposition.

4. Conclusions According to the results of the present study, there is an indication that supplementation with hydrolysable tannins reduces carcass fat deposition, leading also to a higher percentage of PUFA in fat tissue. With regard to meat quality, the results on fresh meat (increased markers of lipid and protein oxidation accompanied by reduced water holding capacity) are indicative of increased prooxidative potential. Supplementary data to this article can be found online at http://dx. doi.org/10.1016/j.meatsci.2017.06.012.

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