Deionised water was purified with a Milli-Q^® water system (Millipore, Bedford, MA, USA). Acetonitrile and methanol were of high-performance liquid chromatography (HPLC) grade and purchased from Sigma Aldrich (Saint Louis, USA), as were the following: (+)-catechin, malvidine-3-O-glucoside, gallic acid, Folin–Ciocalteu’s phenol (2N), sodium metabisulfite, sodium carbonate, phloroglucinol, L-ascorbic acid, hydrochloric acid (37 %), formic acid (98 %), 2,2’azobis(2-methylpropionamidine) dihydrochloride (AAPH, 97 %), disodium phosphate, sodium dihydrophosphate, fluorescein, trolox, polyvinylpolypyrrolidone (PVPP, 110 µm), sodium acetate, syringaldazine, 2,2-diphenyl-1-picrylhydrazyl (DPPH), sodium hydroxide and acetic acid.

Mature grapes (Vitis vinifera, cv. Merlot noir) were collected from several Saint-Émilion vineyards of the Union de Producteurs de Saint-Émilion (Saint-Émilion, south-west France) in 2017. The vineyards had been planted on sandy-gravely or chalky-clay soils with a density of 5,500 vines/ha. The vineyards had not been sprayed with anti-Botrytis synthetic fungicides during the season.

Healthy grape bunches (approximately 66 kg) were randomly selected and then inoculated as described hereafter. The inoculum originated from a 7-day-old culture in Petri dishes of the B. cinerea strain 213. This single-spore isolate was selected from a collection belonging to UMR SAVE, Bordeaux, since it is a highly virulent strain belonging to the transposa type (Martinez et al., 2003; Martinez et al., 2005; Martinez et al., 2008). Fruits were processed in 2-kg batches of grape berries. They were inoculated until the complete fruit surface was covered by spraying a mycelio-spore suspension at the rate of 1.4 x 10⁵ propagules/mL, with fungal propagules comprising a third of conidia and two thirds of mycelium fragments about 60-200 µm in length. To promote grey mould development, inoculated grape bunches were incubated in the dark at 13 °C for 16 days in a climatic phytotronic CONVIRON chamber (at 95-100 % relative humidity). Visual sorting of berries was then performed to remove undesirable rotten berries following other fungal development (e.g., Penicillium spp., identifiable via a blue-green colour). Botrytised grapes were used for micro-winemaking.

Typical infected and symptomatic berries (rotten berries often totally covered by Botrytis mycelium and/or sporulation) were added to healthy berries with a proportion of either 20 or 50 % (ratios were based on number of infected berries rather than on their weight), thus ending up with 7-kg batches of grapes. A control with 100 % healthy grapes was also prepared. Every batch was mechanically crushed and destemmed, then collected in a 10-L aluminium tank. A 3 g/hL dose of SO₂ was added to the control tanks only (botrytisation rate 0 %) using a Bisulfite18 solution at 183 ± 3 g/L of SO₂ (Laffort, Bordeaux, France). Commercial tannins from Laffort and IOC were chosen for their various botanical origins and structures: grape seed (Gse), grape skin (Gsk), quebracho (Q), oak (O) and nutgall (N). In the tanks with botrytised grapes, tannins were added at 100 g/hL, or not at all to obtain controls (20 % and 50 % Botrytis controls). The various treatments are presented in Figure 1. There were 13 tanks (1 x 2 with a botrytisation rate of 0 %, 6 x 2 with a botrytisation rate of 20 % and 6 x 2 with a botrytisation rate of 50 %) in duplicate with about 5 L of musts in each.

Saccharomyces cerevisiae yeasts (Zymaflore^®FX10, Laffort) were prepared by rehydration in warm water (37 °C) for 15 minutes. Rehydrated yeasts were applied to each fermentation tank in order to reach concentrations of 20 g/hL. An alcoholic fermentation activator (Thiazote^®, Laffort) was added along with the leaven when the nitrogen concentration was lower than 140 mg/L and during fermentation (when a density of 1.06 kg/L had been reached) to complete nutriments at 200 mg/L. Fermentations were conducted at a constant temperature of 25 °C and temperature and density were monitored daily in each tank. The cap was punched down daily and O₂ (2 to 4 mg/L) was added when the density had reached 1.02. When the density was stable (0.993-0.997) in each batch and concentrations of reducing sugars were lower than 2 g/L, a 2-day post-fermentation maceration was conducted at 30 °C under CO₂.

Table

(Ø = no addition of tannins; Gse = grape seed tannins; Gsk = grape skin tannins; O = oak tannins; Q = quebracho tannins; N = nutgall tannins; AF = alcoholic fermentation; MLF = malolactic fermentation; VAT = vatting; AF1, AF2, AF3 = 1, 2, 3 days after beginning of alcoholic fermentation respectively; MAC = after post-alcoholic fermentation maceration; WINE = 3 months after bottling).

Lightness (L*), position in a red-green axis (a*) and position in a yellow-blue axis (b*) (CIE, 1986) were measured with a Konica Minolta CM-5 apparatus (Nieuwegein, Netherlands). C*_ab and h_ab were calculated as follows:

Table

Absorbances at 420, 520 and 620 nm were measured in a 1-mm path length cuvette by a UV-Vis Helios Alpha^TM spectrophotometer (Thermo Fisher Scientific Inc., Waltman, MA, USA). Chromatic parameters (Glories, 1984) were calculated as follows: were calculated as follows:

Table

Samples were previously diluted with a dilution factor (DF) of 50 (VAT-samples) or 100. The absorbance at 280 nm was measured in a 1-cm path length quartz cuvette and TPI was calculated as (Ribéreau-Gayon et al., 2006):

Table

FCI was determined according to Singleton and Rossi (1965) with the following solutions placed in 50-mL volumetric flasks: 2.5 mL of Folin–Ciocalteu solution, 0.5 mL of sample or standard solution, 10 mL of Na₂CO₃ solution and water up to the 50-mL mark. After 30 min, absorbance was measured at 760 nm on a UV-vis spectrophotometer. Gallic acid was used as standard at concentrations ranging from 100 to 800 mg/L in order to express FCI in mg of gallic acid equivalents per litre of must or wine.

Total anthocyanins were determined according to Ribéreau-Gayon and Stonestreet (1965). Solution A containing 250 μL of sample, 250 μL of ethanol acidified with 0.1 % HCl and 5 mL of 2 % HCl (v/v) was prepared beforehand. In two tubes, 1 mL of solution A was added to 400 μL of distilled water in tube 1 and 400 μL of 15 % potassium bisulfite in tube 2. After 20 min, the absorbance at 520 nm from both tubes was measured in a 1-cm optical path cuvette. Malvidin-3-O-glucoside was used as standard and TA was expressed in mg/L^{malvidin-3-O-glucoside equivalent.}

All solutions were prepared in 7.4 pH phosphate buffer at 37 °C. In a 96-well black plate, 30 μL of diluted musts or wines (1/2000) or trolox (2.5 to 40 μM), 180 μL of 117 nM fluorescein and 90 μL of 40 mM AAPH solution were added to each well. The fluorescence (excitation = 485 nm and emission = 520 nm) was measured every minute for 90 min at 37 °C. For each kinetic, the area under the curve (AUC) was calculated and then from the standard curve AUC = f (trolox concentration), the antioxidant capacity of each sample was determined as being the concentration mM trolox equivalent (González-Centeno et al., 2012).

According to Milat et al. (2019), 190 μL of 60 μM DPPH in methanol and 10 μL of the diluted sample (1/25 for the wines and 1/10 for the musts) or solutions of trolox (0.1-1 mM) were deposited in a 96-well transparent plate. The absorbance at 515 nm of each well was measured at 25 °C after 30 min. The result was expressed in mM trolox equivalent.

Musts and wines were filtered (0.45 µm) and directly injected for HPLC analyses. HPLC-UV analyses were performed by means of a Thermo Scientific Accela (Thermo Fisher Scientific, Waltham, MA, USA) with an Accela 600 pump module and a UV-Visible diode array detector and Xcalibur Software according to González-Centeno et al. (2017). Separation was performed on a reverse-phase C18 Nucleosil column (250 x 4.6 mm, 5 µm). The injected volume was 20 µL. The mobile phases were water/formic acid (95:5, v/v) (solvent A) and acetonitrile/formic acid (95:5, v/v) (solvent B), at a flow rate of 1 mL/min. Initial solvent B was set at 10 %. The mobile phase gradient was as follows: 35 % B at 25 min, 100 % B at 35 min, 100 % B from 35 to 40 min, 10 % B at 41 min, and then 10 % B for 4 min before the next injection. Eluting peaks were monitored at 520 nm. The mean peaks were identified by comparison with injected external standards and previous results (Chira, 2009). Concentrations were expressed as milligrams of malvidin 3-O-glucoside equivalents per litre.

Musts and wines were filtered (0.45 µm) and directly injected for HPLC analyses. The equipment used for HPLC analysis consisted of a Thermo-Finnigan UV-Visible detector (UV-vis 200), a Thermo-Finnigan autosampler and a Thermo-Finnigan (San Jose, CA, USA) ternary pump coupled to a Xcalibur data treatment system. Separation was performed on a reverse-phase Lichrosphere 100-RP18 (250 mm x 2 mm, 5 µm; Merck, France) column according to González-Centeno et al. (2012). The mobile phases were water-acidified with formic acid 0.5 % (solvent A) and acetonitrile-acidified with formic acid 0.5 % (solvent B). The gradient at a flow rate of 1 mL/min was 3 % B for 3 min, 3 to 5 % B for 11 min, 5 to 10 % B for 8 min, 10 to 14 % B for 4 min, 14 to 25 % B for 14 min, 25 to 100 % B for 1 min, 100 % B for 7 min, 100 to 3 % B for 2 min, and 3 % B for 5 min. Eluting peaks were monitored by a UV-detector at 280 nm and a fluo-detector (λ_excitation = 280 nm, λ_emission = 320nm). Identification of mean peaks was performed by comparison with injected external standards and previous results (Chira, 2009). Calibration curves were established using the catechin as external standard. The results were expressed as milligrams of catechin equivalents per litre.

Proanthocyanidin mDP was determined for wines by phloroglucinolysis (Drinkine et al., 2007). The oligomeric and polymeric products were depolymerised in the presence of a nucleophilic agent (phloroglucinol) in an acid medium. Briefly, wine (5 mL) was evaporated to dryness using a rotary evaporator and then resuspended in 10 mL of milliQ water. The 10 mL sample (or 5 mL of musts) was passed through a pre-activated C18-SPE column (60 mL) and then washed with 60 mL of milliQ water. The samples were then eluted with 60 mL of methanol and the eluate was dried using a rotary evaporator and resuspended in 2 mL of methanol (extract 1). In an Eppendorf tube, 100 μL of extract 1 and 100 μL of methanol solution containing phloroglucinol at 50 g/L, ascorbic acid at 10 g/L and 0.1 N HCl were added. The tube was placed for 20 min in an oven at 50 °C, after which 1 mL of a 40-mM sodium acetate solution was added. The new extract obtained was filtered onto a 0.45-μm pore diameter syringe filter before injection. Reaction products were analysed by HPLC-MS on an Accela series (Thermo-Scientific, Illkirch-Graffenstaden, France), equipped with a pump module and a UV detector. Separation was performed on a reverse-phase Waters XTerra^®RP C18 (Waters, Milford, MA, USA) (100 mm x 4.6 mm, 3.5 µm) column at room temperature. A binary gradient system was employed using 1 % (v/v) aqueous acetic acid (solvent A) and MeOH (solvent B) at a flow rate of 1 mL / min. The elution conditions were as follows: 5 % B for 25 min; 5-20 % B for 20 min, 20-32 % B for 15 min, and 32-100 % B for 2 min. The column was then washed with 100 % B for 5 min and re-equilibrated with 5 % B for 5 min before the next injection. The mDP, % of prodelphinidin (% P) and % of galloylation (% G) were estimated using the response factors of proanthocyanidin cleavage products at 280 nm and calculated as described by Chira et al. (2012). The mDP was obtained by dividing the sum of all subunits (flavan-3-ol monomers and phloroglucinol adducts, expressed in moles) by the sum of all flavan-3-ol monomers (expressed in moles).

Several triangular tests (ISO4120: 2007) were carried out to determine significant differences between

i) the main control wine [0 % Botrytis, Ø tannins] and the other control wines [20 % Botrytis and 50 %, Ø tannins], and

ii) between control wines [20 % and 50 % Botrytis, Ø tannins] and the corresponding wines supplemented with tannins.

During the sessions, the judges were asked to observe, smell and/or taste the samples in the defined order and indicate which one seemed different, even if they were not sure of their answer. The panel was also asked to freely note one or more descriptor(s) in order to differentiate the samples. The descriptors were retained only in the event of a correct answer from the judge.

Ranking tests (ISO8587: 2007) were conducted in two sessions of six wines each (for each botrytisation rate). The panel of tasters classified the six tested wines according to the "overall quality" of the product (visual, aromatic and taste qualities), knowing that they were young wines (three-months old) made from Merlot. Ratings were assigned from 1 (the most qualitative) to 6 (the least qualitative). Results were processed using a Friedman test on the sum of the ranks assigned by the judges. The smallest significant difference was then calculated and compared to the absolute value of the difference in the sum of the ranks (for two wines), to determine which products were significantly different from each other.

The wines were obtained from healthy grapes (0 % Botrytis) and from a mix of healthy grapes and totally infected grapes to reach a final 20 % or 50 % botrytisation rate. Table 1 presents the oenological and chemical parameters of the control (0 %), 20 % and 50 % botrytised musts and their corresponding 3-month-old wines.

The classic oenological parameters (pH, density, concentrations of reducing sugars and total sugars) varied slightly according to the level of botrytisation of the musts, although significant differences were found. Volatile acidity content increased by 17 % with a botrytisation level (trend) of 0 to 20 %, and significantly by 36 % with a 20 % to 50 % botrytisation level. Beside the possible production of acetic acid by Botrytis, the highest volatile acidity at higher botrytisation levels can also be explained by the action of acetic bacteria, most likely present in damaged grapes (Ribéreau-Gayon et al., 2012).

The greatest differences were observed for phenolic compounds. The total phenolic compounds in botrytised musts decreased significantly to levels approximately two-fold less significant (TPI) or even three-fold less significant (FCI) than in the healthy musts. Anthocyanins were also drastically degraded in both 20 % and 50 % botrytised musts. Their concentration decreased significantly from 108.2 (control must) to less than 10 mg/L malvidin-3-O-glucoside eq. in the botrytised musts (anthocyanins identified and quantified by HPLC in botrytised musts in trace amounts). These results may be due to the presence of laccase with corresponding activities of 25.8 LU and 42.9 LU in 20 % and 50 % botrytised musts respectively (Table 1). Hence, there was sufficient laccase activity in the 20 % botrytised must to degrade all the anthocyanin content in the healthy musts. The major anthocyanin compound, malvidin-3-O-glucoside, present in the healthy must at about 35 mg/L, was thus nearly absent from both botrytised musts. These results are in agreement with those obtained by Ky et al. (2012). Moreover, the results in Table 1 show that the molecular tannins were significantly affected by enzymatic oxidation. Like the anthocyanins, the tannins had probably been oxidised since they are also substrates of laccases (Oliveira et al., 2011).

Table

titre du tableau
	Musts	3-month-old wines
% Botrytis	0 %	20 %	50 %	0 %	20 %	50 %
Oenological parameters
pH	4.8 a*	± 0.0	4.8 a	± 0.0	4.7 b	± 0.0	3.7	± 0.0	3.8	± 0.0	3.8	± 0.0
Titratable acidity (g/L H2SO4 eq)	3.4	± 0.0	3.7	± 0.2	3.7	± 0.3	3.5	± 0.1	3.2	± 0.1	3.6	± 0.2
Density (g/dm3)	1105 a	± 0	1104 b	± 0	1106 c	± 0	991 a	± 0	993 ab	± 0	995 b	± 1
Alcohol (% v/v)	-	-	-	12.8 a	± 0.1	12.2 b	± 0.0	12.3 b	± 0.0
Volatile acidity (g/L H2SO4 eq.)	0.45 a	± 0.00	0.53 a	± 0.00	0.72 b	± 0.04	0.34 a	± 0.01	0.52 b	± 0.02	0.90 c	± 0.20
Sugars
Reducing sugars (g/L)	167.4 a	± 0.0	162.8 b	± 0.4	164.6 ab	± 1.3	1.6	± 0.1	1.4	± 0.1	1.6	± 0.3
Total sugars (g/L)	194.4 a	± 0.0	192.2 b	± 0.1	197.8 c	± 0.0	6.2 a	± 0.3	7.7 ab	± 0.2	10.2 b	± 1.1
Organic acids
Tartaric acid (g/L)	3.8 a	± 0.0	3.4 a	± 0.0	2.6 b	± 0.2	2.0	± 0.1	2.2	± 0.0	2.4	± 0.3
Malic acid (g/L)	2.0 a	± 0.0	2.8 b	± 0.1	2.6 b	± 0.1	0.4	± 0.0	0.2	± 0.0	0.3	± 0.0
Lactic acid (g/L)	-	-	-	1.9 a	± 0.0	1.7 b	± 0.0	1.5 c	± 0.0
Phenolic compounds
TPI	12.4 a	± 0.7	5.4 b	± 0.3	7.6 b	± 0.7	45.8 a	± 1.5	43.9 a	± 0.6	37.4 b	± 1.7
FCI (mg/L gallic acid eq.)	436 a	± 1	137 b	± 10	186 b	± 25	1932 a	± 62	1692 b	± 45	1424 c	± 59
TA (mg/L malividin-3-O-glucoside eq.)	155.1 a	± 0.6	0.2 b	± 0.2	4.9 c	± 0.2	547.4 a	± 1.1	347.6 b	± 36.4	124.3 c	± 16.6
Molecular anthocyanins (mg/L malividin-3-O-glucoside eq.)
delphinidin-3-O-glucoside	8.5	± 0.4	< 2	< 2	29.5 a	± 0.9	14.1 b	± 2.5	5.3 c	± 0.1
cyanidin-3-O-glucoside	10.4	± 0.3	< 2	< 2	8.6 a	± 0.6	7.1 a	± 0.2	4.7 b	± 0.0
petunidin-3-O-glucoside	7.8	± 0.2	< 2	< 2	31.5 a	± 0.0	17.8 b	± 2.5	7.3 c	± 0.6
peonidin-3-O-glucoside	23.2	± 0.6	< 2	< 2	30.9 a	± 1.2	21.9 b	± 0.1	10.5 c	± 0.4
malvidin-3-O-glucoside	34.9	± 0.6	< 2	< 2	128.6 a	± 3.3	85.3 b	± 8.4	31.6 c	± 5.0
peonidin-3-O-(6-O-acetyl)-glucoside	5.5	± 0.0	-	-	12.9 a	± 0.5	8.1 b	± 0.7	4.8 c	± 0.2
malvidin-3-O-(6-O-acetyl)-glucoside	10.7	± 0.5	-	-	40.0 a	± 0.3	21.7 b	± 3.8	8.1 c	± 0.6
peonidin-3-O-(6-O-p-coumaroyl)-glucoside	3.0	± 0.1	-	-	8.0 a	± 0.3	6.0 b	± 0.3	4.2 c	± 0.1
malvidin-3-O-(6-O-p-coumaroyl)-glucoside	4.2	± 0.3	-	-	16.7 a	± 0.1	10.7 b	± 1.6	5.2 c	± 0.3
Total anthocyanins identified	108.2 a	± 1.4	< 10.0 b	± 0.0	< 10.0 b	± 0.0	306.6 a	± 1.9	192.6 b	± 19.9	81.5 c	± 7.3
Molecular tannins (mg/L catechin eq.)
B1 (dimer)	0.01	± 0.0	-	-	0.9	± 0.1	-	-
B3 (dimer)	4.6	± 0.4	-	-	17.0	± 6.5	5.8	± 1.6	0.6	± 0.5
catechin	10.0	± 0.7	< 2	< 2	68.4 a	± 1.9	42.6 b	± 4.6	5.1 c	± 2.9
B4 (dimer)	0.7	± 0.1	-	-	5.7 a	± 1.1	0.8 b	± 0.5	0.3 b	± 0.0
B2 (dimer)	1.4	± 0.1	-	-	10.2 a	± 1.2	3.6 b	± 1.0	0.8 b	± 0.5
epicatechin	4.9	± 0.1	-	-	54.0 a	± 0.8	19.2 b	± 8.8	1.7 b	± 1.3
T (trimer)	17.7 a	± 0.7	3.8 b	± 0.0	3.7 b	± 0.1	13.3 a	± 0.1	6.9 ab	± 0.8	4.0 b	± 2.8
Total tannins identified	39.4 a	± 1.8	3.9 b	± 0.0	3.8 b	± 0.1	169.5 a	± 3.1	78.9 b	± 17.3	12.4 c	± 7.9
mDP	-	-	-	3.2 ab	± 0.1	3.5 a	± 0.1	2.8 b	± 0.2
% G	-	-	-	22.6	± 0.3	27.2	± 0.5	22.6	± 4.6
% P	-	-	-	16.2 a	± 0.4	9.0 b	± 0.3	8.9 b	± 2.1
% C	-	-	-	61.3 a	± 0.7	63.8 ab	± 0.3	68.6 b	± 2.5
Antioxidant capacity
ORAC (mM eq. Trolox)	5.0	± 0.3	-	-	59.2	± 3.2	57.6	± 0.7	51.1	± 0.7
DPPH (mM eq. Trolox)	2.7 a	± 0.1	0.4 b	±0.4	-	12.9	± 0.9	9.9	± 0.1	8.3	± 1.8
Spectrophotometric parameters of colour
CI	0.7	± 0.3	0.7	± 0.0	0.8	± 0.1	9.3 a	± 0.2	8.0 b	± 0.0	5.5 c	± 0.0
Hue	1.1 a	± 0.2	2.6 b	± 0.0	2.6 b	± 0.0	0.6 a	± 00	0.8 a	± 0.1	1.2 b	± 0.0
% yellow	47.5 a	± 3.7	65.6 b	± 0.0	65.6 b	± 0.2	32.1 a	± 0.1	38.0 b	± 2.0	48.8 c	± 0.7
% red	44.8 a	± 3.3	25.7 b	± 0.3	25.4 b	± 0.7	56.7 a	± 0.0	50.5 b	± 2.1	41.0 c	± 0.6
% purple	7.6	± 0.5	8.7	± 0.3	9.0	± 0.0	11.1 a	± 0.1	11.5 a	± 0.1	10.2 b	± 0.1
CIELAB parameters
L*	79.0	± 5.7	86.9	± 0.6	83.2	± 1.4	14.6 a	± 0.1	15.3 a	± 0.3	27.0 b	± 0.0
a*	18.8	± 9.4	3.1	± 0.5	6.5	± 1.7	46.2 a	± 0.0	45.1 a	± 0.4	50.9 b	± 0.3
b*	15.2 a	± 0.4	24.7 b	± 1.2	29.2 c	± 0.4	24.6 a	± 0.1	25.2 a	± 0.0	39.3 b	± 0.8
Ca.b	24.6 a	± 7.0	24.9 b	± 1.2	29.9 c	± 0.7	52.3 a	± 0.0	51.6 b	± 0.3	64.3 c	± 0.2
ha.b	41.0 a	± 15.1	82.9 b	± 0.7	77.5 ab	± 3.0	28.0 a	± 0.1	29.2 a	± 0.2	37.7 b	± 0.7
Laccase activity
Laccase activity nmol/(min*mL)	-	25.8 a	± 0.4	42.9 b	± 2.0	-	0.5 a	± 0.1	6.9 b	± 1.0

* Different letters show significant differences between different botrytisation levels for the same variable (p-values < 0.05); a lack of letter indicates no significant difference. TPI = total polyphenol index, FCI = Folin-Ciocalteu index, TA = total anthocyanins, mDP = mean degree of polymerisation, % G = % of galloylation, % P = % of prodelphinidins, % C = % of procyanidins, CI = colour intensity.

Furthermore, since the chromatic characteristics and antioxidant capacities of botrytised musts were also markedly affected, the consequences of the degradation of phenolic compounds by laccase were again directly observable (Table 1); for example, the spectrophotometric “hue” parameter (A_{420 nm} / A_{520 nm}) was approximately twice as high in botrytised musts as in healthy musts, indicating a major oxidative degradation of botrytised musts.

The presence of laccase produced by B. cinerea thus had a considerable negative impact on the phenolic compounds of the musts as soon as they were in the vat, the degradation having begun in the grape berries (Ky et al., 2012). The oxidation of polyphenols by laccase therefore directly affected the visual characteristics and antioxidant properties of the musts, which is expected to impact the final quality of the wines.

In contrast to the wines from healthy Merlot grapes, the botrytised control wines obtained from the botrytised musts were damaged, their colour and phenolic compounds having been drastically affected (Table 1). The botrytised wines were less intense (CI); they had a stronger yellow hue (increasing % yellow and decreasing % red) and a brickish colour. These characteristics increased with botrytisation rate. Tannin and anthocyanin contents decreased with botrytisation level.

All wine variables (oenological, chromatic, spectrophotometric, fluorometric and chromatographic parameters of the phenolic compounds and laccase activity) are represented in a PCA plot (Figure 2), with three main axes accounting for over 83 % of the overall variability.

In the PCA plot, the first PCA axis classifies the wines according to their botrytisation rate very clearly (increasing from right to left). However, two wines were differentiated owing to positive axis-2 coordinates; i.e., the wines at 20 % and 50 % botrytisation levels with added nutgall tannins (N). These wines had higher TPI, ORAC and DPPH values than the other wines. The plot of 1 and 3 axes shows differences between wines supplemented with the various tannins, especially within the 20 % botrytisation group. Wines with Gse and Gsk tannins were the furthest from the control wines without added tannins, suggesting that their addition had a positive impact on the final phenolic content (Figure 2B). Wines obtained from 50 % botrytised grapes were more grouped, showing lower variability. As a preliminary result, the added oenological tannins at 100 g/hL may have had less effect on the studied variables in the 50 % botrytised wines than in the 20 % botrytised wines.

Table

A) plot of axes 1 and 2, B) plot of axes 1 and 3. Gse = grape seed tannins, Gsk = grape skin tannins, Q = quebracho tannins, O = oak tannins, N = nutgall tannins. Groups corresponding to the level of botrytisation are highlighted in green (50 %), red (20 %) and black (0 %).