America n Journal of Analy tic al Chemistry, 2011, 2, 344-351
doi:10.4236/ajac.2011.23042 Published Online July 2011 (
Copyright © 2011 SciRes. AJAC
Use of Ultrasound Bath in the Extraction and
Quantification of Ester-Linked Phenolic Acids in Tropical
Mellina Damasceno Rachid Santos1, Aline de Paula Vitor1, Jailton da Costa Carneiro2,
Domingos Sávio Campos Paciullo2, Renato Camargo Matos1, Maria Auxiliadora Costa Matos1*
1NUPIS (Núcleo de Pesquisa em Instrumentação e Separação Analíticas), Departamento de Química, Instituto de
Ciências Exatas, Universidade Federal de Juiz de Fora, Juiz de Fora, Brazil
2EMBRAPACentro Nacional de Pesquisa de Gado de Leite (CNPGL), Juiz de Fora, Brazil
Received March 29, 2011; revised May 12, 2011; accepted May 20, 2011
A method was developed for the analysis of ester-linked phenolic acids in forage samples using extraction by
an ultrasound-assisted treatment and quantification by HPLC with a UV -VIS detector. A reversed-phase C18
column was used for developing the method and the optimal condition was established with isocr atic eluti on
using acetonitrile/methanol/H3PO4 pH 2.08 (13:12.5:74.5) as the mobile phase. To reduce the time of sample
processing, the extraction of est er-linked phenolic acids was studied using ultrasound bath and the results
were then compared with those from an extraction usual using alkaline hydrolys is ( 20˚C for 24 h). The me-
thod was valued through external and internal calibration. Internal calibration using o-coumaric acid as in-
ternal standard and m-coumaric acid as surrogate internal standard showed better results. The detection limits
were of 0.09 and 0.04 mg·L–1 for p -coumaric and ferulic acids, respectively. The proposed method showed a
good linear dynamic range (3.00 - 30.00 mg·L–1) for the analytes. The usefulness of the method- ology was
demonstrated by addition-recovery experiments using forage samples and values were in the 83 to 99%
range. The extraction of ester-linked phenolic acids by 120 minutes of ultrasound bath was faster and more
reproducible than alkaline hydrolysis (20˚C for 24 h).
Keywords: Ultrasound Bath, Phenolic Acids, Forage, Internal Calibration
1. Introduction
Phenolic compounds are a group of secondary metabo-
lites synthesized by plants during development as a de-
fense against pathogenic ingress, wounding and UV ra-
diation [1,2]. Among the main phe nolic compounds pre-
sent in plants are phenolic acids, such as ferulic acid, p-
coumaric acid and caffeic acid.
These acids are present in plants in both free and
bound forms where a small fraction occurs as “free ac-
ids” and the majo rity are linked to structural components
of the plant [3,4]. According to Jung (1989), ferulic and
p-coumaric acids seem to be generally esterified to dif-
ferent components of the cell wall. The majority of p-
coumaric acid are esterified to lignin whereas ferulic acid
tends to associate with the hemicellulose fraction and
may form a cross-linka ge between li gnin and hemicellu-
lose [3,4]. Due t o such asso ciati ons, seve ral stud ies ha ve
shown that these compounds potentially affect the di-
gestibility of forages by the rumen, thereby compromis-
ing li vestock p erformance [3,5-10].
Acidic hydrolysis and alkaline hydrolysis are the most
common means of releasing the phenolics acids [2]. The
main method of extraction applied in forage samples is
alkaline hydrolysis. It allows quantification of total phe-
nolic acids (ester-linked and ether-linked) or those that
are only ester-linked through two types of treatment:
mild alkaline or hot alkaline hydrolysis [11]. In the mild
alkaline extraction, where the sample is treated with
NaOH 1 mol·L–1 solution at 20˚C for 24 h, only ester
bonds are cleaved. During hot alkaline hydrolysis, the
sample is usually treated with a solution of NaOH 4
mol ·L–1 at 170˚C for 2 h, and both ester and ether bonds
are cleaved. The ether-linked derivatives are estimated
Copyright © 2011 SciRes. AJAC
by the difference between total phenolic acids and the
ester-linked molecules [4,7].
Phenolic compounds have been analyzed in different
samples by capillary electrophoresis (CE) [12-15], gas
chromatography equipped with a mass spectrometric
detector (GC-MS) [16-20], ultra performance liquid
chromatography (UPLC) [21] and high performance liq-
uid chromatography (HPLC) [3,5-7,9,20-27]. HPLC is
most frequently used because it does not require a deri-
vatization sample for analysis as gas chromatography
[24]. When compared to c apillar y electrophoresis, HP LC
is better in terms of accuracy, sensitivity and precision
although it consumes more solvent and time for sample
treatment [15].
Recently, ultrasound bath has been used as an ancil-
lary tool for traditional extraction methods due to the
simplicity of the method and a decrease in the time
needed for extraction. There have been many studies
applying ultrasound bath in the sample treat ment of soil s
and sediments [28-31], biological [32-34 ], nut ritio us [35,
36]. However, in the case of forage samples, no study
has yet been reported in the literature describing this
technique for the extraction and quantification of pheno-
lic acids.
The objective of this work was to propose an a lter na-
tive method that is fast and precise for extracting ester-
linked phenolic acids using an ultrasound bath. This pa-
per describes the development of a new method using
high performance liquid chromatography (HPLC-UV)
for the separation of five phenolic acids (ferulic, p-cou-
maric, m-coumaric, o-coumaric and caffeic acids). Shiki-
mic acid was also included in the calibrations and is an
organic acid that occurs as an intermediate during the
process of hydroxycinnamic acid formation [3,37]. The
performance of the analytical procedure was evaluated
by determining the concentration of phenolic acids in
forage samples.
2. Experimental
2.1. Chemicals and Solutions
The standard compounds (including ferulic, p-coumaric,
m-coumaric, o-coumaric, caffeic and shikimic acids)
were purchased from Sigma-Aldrich (St. Louis, MO,
USA). HPLC grade solvents (methanol, acetonitrile and
tetrahydrofuran) and HPLC grade reagents (phosphoric
acid, acetic acid and potassium phosphate monobasic)
were from Tedia Company Inc. (Fairfield, OH, USA).
The chemicals sodium acetate, sodium hydroxide and
hydrochloric acid were from Vetec (Rio de Janeiro, RJ,
Stock solutions of the ferulic, p-coumaric,
m-coumaric, o-coumaric, caffeic and shikimic acids were
prepared at a concentration of 1 g·L–1 in methanol and
filtered thr- ough a 0.45 µm PT FE filter. These solutions
were stable for one week whe n stored at 4˚C.
2.2. Preparation of Calibration Standards
Standard solution containing a mixture of caffeic, shiki-
mic, ferulic and p-coumaric acids were first prepared in
mobile phase at a concentration of 100 mg L-1 and were
gradually diluted in mobile fase to working concentra-
tions of 3 to 20 mg·L–1. An internal standard of o-cou-
maric acid and a surrogate internal standard of m-cou-
maric acid was added to set up a resulting concentration
of 10 mg·L–1 in all standard solutio ns.
2.3. HPLC Apparatus
HPLC analyses were conducted using an Agilent 1100
Series system equipped with a manual injection valve
with a 20 µL sample loop, a degasser system, a quartet
pump and a multiple wavelength UV-detector. Data ac-
quisition and processing were accomplished with the
Agilent Chemistation LC Systems software. A reversed-
phase (RP) column ZORBAX ODS (150.0 mm × 4.6
mm I.D., 5 μm particle size) and a ZORBAX ODS
pre-column (12.5 mm × 4.6 mm I.D., 5 μm p article size)
were used at room temperature.
2.4. Optimization of the Chromatographic
This study involved the optimization of the separation
conditions of shikimic, caffeic, p-coumaric and ferulic
acids using a mobile phase composed for organic sol-
vents and acid solution or buffer. Initially, mixtures of
methanol, acetonitrile or tetrahydrofuran with aqueous
solution were tested as eluents to obtain a sufficient
resolution. The pH range of the eluent (2.08 to 4.50) was
adjusted with phosphoric acid, acetic acid, acetate buffer
or phosphate buffer. The values of retention time, peak
symmetry, resolution and sensitivity were evaluated for
sufficient reso lution on the RP- column.
2.5. Plant Material
Fractions of stem and leaf of the species Brachiaria bri-
zantha Marandu, Cynodon dactylon Florakirk, Cynodon
nlemfuensis Florona, Panicum maximum Mombaça,
Pennisetum purpureum Anão and Pennisetum pur- pu-
reum P ioneiro were obtained from the experimental farm
of EMBRAPA (Goiás, Brazil). The samples were taken
every 30 days, dried in forced air oven at 55˚C for
Copyright © 2011 SciRes. AJAC
Table 1. Content (mg·g1 dry matter) of ester-linked p-cou maric and ferulic acids in Panicum maximum Mo mbaça ext racted
in bath with pr o grammed t emp erat ure (a t 20˚C for 2 4 h) and ultrasound (at room temperature for 120 min).
Content (mg·g–1 dry matter)
Bath with programmed temperature Ultrasound
Rep l icate p-CA FA p-CA FA
1 4.48 4.75 4.08 4.93
2 4.39 4.52 4.09 4.93
3 4.20 4.42 4.05 4.88
Average 4.35 4.56 4.07 4.91
Sd 0.14 0.17 0.03 0.03
RSD 3 3 1 1
p-CA = Ester-Linked p-Coum aric A cid; FA = Ester-Linked Feru lic Acid; Sd = standard deviation; RSD = relative standard deviation.
72 h and grounded (1 mm).
2.6. Extraction of Ester-Linked Phenolic Acids
Approximately 25.0 mg of leaf of Panicum maximum
Mombaça were extracted at room temperature with 1
mol ·L–1 NaOH using the following times for sonication:
15, 30, 60, 90, 120, 150, 180, 210, 240, 270 and 300 mi-
nutes. The results obtained were then compared with
those fro m the al kal ine hydr olys is a t 20˚C for 24 h. Trip-
licate extractions and analyses were carried out.
After extraction, the samples were subsequently fil-
tered and washed with water. The combined filtrate and
wash was acidified to pH 2.5 with 6 mol ·L–1 HCl and
brought to a final volume of 10 mL. The sample solu-
tions were diluted using mobile phase, filtered through a
0.45 µm PTFE filter and analyzed by HPLC. The injec-
tion volume was 20 µL.
3. Results and Discussion
3.1. Development of the HPLC Me tho d
Preliminary tests employing a binary mixture (organic
solvent: aqueous solution) as the mobile phase were per-
formed. T he best isocratic separation was established for
each organic solvent (acetonitrile, methanol and tetrah y-
drofuran) at different pH values (2.08, 2.20, 2.50, 3.00,
3.50, 4.00 and 4.55). This showed that increasing pH
resulted in a longer retention time of the compounds.
Depending on the eluent pH, the ferulic and p-coumaric
acids present double peaks at higher pH value s. This c an
be explained by the presence of weak organic acids, wit h
pKa values of around 4.5 and 9.5 [17]. The best results
were obtained with the mobile phase adjusted to pH 2.08
with phosphoric acid (for all three solvents) because
ionization of the phenolic acids is suppressed at this pH
value. Ho wever, p H var iatio ns of t he el uent us ing b inar y
compositions did not improve the resolution. Amon g the
compositions of the mobile phase optimized by applying
the ternary mixture of solvents, the best separation con-
dition was for isocratic elution with a mobile phase
composed of acetonitrile/methanol/H3PO4 pH 2.08 (13:
12.5:74.5) at a flow of 1 mL·L–1. The detection of the
compounds was based on different wavelengths that took
into consideration their maximum absorption. Here, the
signal was registered at 236 nm from 0 to 3 minutes, 316
nm from 3 to 9.1 minutes and 236 nm from 9.1 to 15
minutes. This detection had increased sensitivity, im-
proving the detection and quantification limits of the
3.2. Extraction of Ester-Linked Phenolic Acids
For ultrasound bath extraction, analyses were performed
in three replicates. Sonication times of 90 minutes and
less were not enough to extract completely the ester-
linked phenolic acids. The peak areas of the phenolic
acids increased with increasing time and reached their
maxima at 120 minutes of sonication. At sonication
times over 150 minutes, the peak areas decreased, indica-
ting a possible compound degradation.
To evaluate accuracy, paired Student's t-test was used
to determine whether significant differences existed be-
twee n results obtained usi ng 120 minutes of sonication in
an ultrasound bath and thos e ob tained by alka line hydro-
lysis extraction (20˚C for 24 h). The paired Student's t-
test (
= 0.05) showed that there was not a significant
difference between the means. However, results from
extraction employing the ultrasound bath presented s-
maller relative standard deviations, indicating that this
method is more reproducible when compared to alkaline
hydrolysis (Table 1). Therefore, extraction with ultra-
sound bath for 120 minutes was used to reduce the time
of sample a nalysis.
3.3. Features of the Analy tical Meth od
The evaluat ion o f the method was p erfor med b y external
Copyright © 2011 SciRes. AJAC
Figure 1. Typical chromatograms of (a) standard mixture
and (b) phenolic acids released from ester bonds in a sam-
ple of Panicum maximum Mombaça leaf. Peaks: 1) shikimic
acid, 2) caffeic acid, 3) p-coumaric acid, 4) ferulic acid, 5)
m-coumaric acid, and (6) o-coumaric acid. Analytical con-
ditions: column, ZORBAX ODS; fl ow-rate, 1.0 ml/min;
detection for programming of wavelength; mobile phase,
acetonitrile/methano l/H3PO4 pH = 2.08 (13:12.5:74.5).
and internal calibration. Initial ly was studie d the applica-
tion of the o-coumaric and m-coumaric acids as internal
stand ard and surrogate internal stand ard . Preli minar y wa s
studied the presence of o-coumaric and m-coumaric ac-
ids in forage samples. Figure 1 compares the chroma-
tograms of a standard mixture (ferulic, p-coumaric, m-
coumaric, o-coumaric, caffeic and shikimic acids) and a
forage sample. The retention time obtained for the com-
pounds confirmed that o-coumaric and m-coumaric acids
were not present in the samples. As a consequence, m-
coumaric and o-coumaric acids were used as surrogate
internal sta nda rd and internal standard, respectively.
The precision of the method was evaluated by re-
peated injection (
= 6) of a forage sample, and the
standard deviation was determined as better than 3%
(external calibratio n) and 1% (internal calibration). The
sensitivity was also determined using the limit of detec-
tion (LOD) and the limit of quantification (LOQ). The
LOD was calculated as 3 s, where s is the average signal
of standard deviation of 6 forage sample injections with
low phenolic acid concentration, and the LOQ was 10 s.
Table 2 shows the analytical parameters for repeated
injection of a tropical forage sample using internal and
external calibrations. The internal calibration presented
smaller values of LOD, LOQ and RSD. The best results
were attributed the use of the internal standard (o-cou-
maric acid) and surrogate standard (m-coumaric acid)
that corrected fluctuations between each injection and
losses during the sample extraction process, respectively.
Linearity was evaluated, taking into account the cor-
relation coefficient (r) and the response factor in the
Table 2. Limit of detection (LOD), limit of quantification
(LOQ ) and re peatability ac hieved f or external and interna l
External c alibration Internal calibration
ound LOD
(mg·L–1) LOQ
(mg·L–1) RSD
(%) LOD
(mg·L–1) LOQ
(mg·L–1) RSD
p-CA 0.18 0.60 3 0.09 0.28 1
FA 0.08 0.26 1 0.04 0.15 1
p-CA = Ester-Linked p-Coumaric Acid; FA = Ester-Linked Ferulic Acid;
LOD = limit of detection; LOQ = limit of quantification; RSD = relative
standard deviati on.
Table 3. Results of regres s ion analys is on cali bration.
External c alibration Internal calibration
equati o n y =
ax + ba
equati o n y =
ax + bb
y = 17.392x +
r = 0.99 940
y = 0.1987x +
r = 0.99 970
y = 111.81x +
r = 0.99 995
y = 1.2778 x
r = 0.99 995
y = 154.71x +
9.4925 r = 0.999 90
y = 1.768 x
0.0026 r = 0.999 90
Ferul ic acid
y = 96.016x +
r = 0.99 910
y = 1.097 3 x
+ 0.039 9
r = 0.99 935
aWhere y and x are the peak area (mAU) and concentration of the analytes
(mg·L–1), respectively; bWhere y is the rati on of the area of the ana lyte peak
divided by the area of the surrogate intern al standard and x is the ration of
the concentration of the analyte divided by the concentration of the surro-
gate internal standard.
concentration range of 3.00 to 30.00 mg·L–1. Calibration
curves were determined by a mixture of standard solu-
tions of t he pheno lic acids, an d applying an internal stan-
dard (o-coumaric acid) and a surrogate standard (m-
coumaric acid). Each point on the calibration curve cor-
responds to an average signal from three independent
peak measurement s for ea ch aci d. The propor tionalit y of
peak area and concentration was confirmed for all the
analytes (correlation coefficient > 0.999), as shown in
Table 3.
The accuracy of the method was evaluated through
recovery assays using tropical forage samples spiked
with a mixture of acid standards comprising ferulic, p-
coumaric, m-coumaric, caffeic and shikimic acids at
three fortification levels: 5.00, 7.50 and 10.00 mg·L–1
(Table 4) with 10.00 mg·L–1 o-coumaric acid as the in-
ternal standard. A blank spike (NaOH 1 mol·L–1) (
5) was also prepared for the extraction tests with a mix-
ture of the phenolic acids at a concentration of 10.00
mg·L–1 (Table 4).
Table 4 shows the results of recovery for p-coumaric,
ferulic and m-coumaric acids using internal and external
calibrations. The extraction method recoveries obtained
for phenolic acids ranged from 82 to 99%, thus confirm-
ing the accuracy of the method for extraction of phenolic
Copyright © 2011 SciRes. AJAC
Table 4 . Rec o ve ry an d relat ive s tan d ar d de vi at i on ac hi ev ed for sa mpl e a n d bl a nk spi ked wi t h a mixt ure of aci d st a nda rd s f or
external and internal calibration.
Compound S ample Levels of spiked concentration (mg·L–1)
= 3 Internal calibration External calibration
Mea n Recovery (%) RSD (% ) Mea n Recov ery (%) RSD (%)
p-Coumaric acid Sample spike
5.00 98 2 85 4
7.50 98 1 82 4
10.00 99 1 86 4
Blank spike 10.00 91 2 85 3
Ferul ic acid Sample spike
5.00 88 2 85 1
7.50 89 2 82 2
10.00 91 2 89 5
Blank spike 10.00 83 1 85 3
m-Coumaric acid Sample spike
5.00 92 4 84 1
7.50 92 1 83 3
10.00 91 1 91 2
Blank spike 10.00 97 3 95 3
RSD = relative standard deviation.
Table 5 . Content (mg·g–1 dry matter) of ester-linked p-cou mari c and fer ulic aci ds in s a mpl e s o f Brachiaria brizantha, Cynodon
dactylon, Cynodon nlemfuensis, Panic um maximum and Pennisetu m purpure um.
± C.I. (mg·g
dry matter)
Brachiaria bri zantha Marandu stem
Cynodo n dactylon cv. Florakirk
7.89 ± 0.21 4.28 ± 0.07
8.60 ± 0.41 4.11 ± 0.21
Cynodo n nlemfuensis Florona
5.30 ± 0.11
4.48 ± 0.09
Panicum maximum Momb aça s tem 7.74 ± 0.33 4.19 ± 0.10
leaf 4.36 ± 0.14 3.35 ± 0.17
Penniset um p ur pureu m Anão
0.84 ± 0.20 1.63 ± 0.27
1.05 ± 0.07
3.34 ± 0.32
Penniset um p ur pureu m Pioneiro stem
p-CA = Ester-linked p-coumaric acid; FA = Ester-linked ferulic acid; aMean ± c o nfidence inte r v a l (
= 0.05) from 3 determinations.
acids. A better accuracy was obtained by internal cali-
bration with RSD < 4% as the external calibration
showed RSD < 5%. However, caffeic acid is unstable
and it could not be detected after the alkaline hydrolysis
in this study, with no signal being detected at the reten-
tion t ime o f t his anal yte ( Su n et a l., 2001). Shikimic acid
was detected but it was not quanti fied in the spiked tr op-
ical forage sample since it presented co-elution with
compo nents p resent in t he samp le. For the quantificatio n
of shikimic acid, it is necessary to adjust the method, so
that the accuracy was not determined for the recovery of
shikimic and caffeic acids.
3.4. Determination of Est er-Linked Phenolic
Under optimum conditions, the ultrasound bath method
was applied to determine p-coumaric and ferulic acid
concentrations in six tropical forage samples (in tripli-
cate), applying an internal standard (o-coumaric acid)
and a surrogate internal standard (m-coumaric acid). Ta-
ble 5 shows the results obtained for p-cumaric and fer-
ulic acids in fractions of stem and leaf of the species
Brachiaria brizantha Marandu, Cynodon dactylon Flo-
rakirk, Cynodon nlemfuensis Florona, Panicum maximum
Mombaça, Pennisetum purpureum Anão and Pennisetum
purpureum Pioneir o. A reasonably good correlation (
= 0.99 – p-coumaric acid and
= 0.98 ferulic acid)
between the ultrasound bath (2 h) and the thermostatic
bath (20˚C for 24 h) was found for extrac- tion. For
p-coumaric acid, the confidence interval for the slope
and intercept are (0.93 ± 0.03) and (0.01 ± 0.11) mg·g–1
dry matter, respectively, for a 95% confidence level. For
the ferulic acid the confidence interval for the slope and
intercept are (1.07 ± 0.04) and (0.01 ± 0.16) mg·g–1 dry
matter, respectively, for a 95 % confidence level. A
Copyright © 2011 SciRes. AJAC
paired Student´s t-test showed that the mean values
exp crit
; 3.52 for p-coumaric acid and 3.38 for ferulic
acid < 4.30,
= 3,
= 0.95) did not significantly
differ. Taking into account these results, no significant
differences between the extraction methods were ob-
served, strongly indicating the absence of systematic
As can be seen in Table 5 for samples of Brachiaria
brizantha Marandu, Cynodon nlemfuensis Florona, Pen-
nisetum purpureum Anão and Pennisetum purpureum
Pioneiro the concentration of ferulic acid is high in the
leaves while the concentration of p-coumaric acid is high
in the stem o f Brachiaria brizantha Mara ndu and Penni-
setum purpureum Pioneiro. For samples of Panicum
maximum Mombaça, both acids are found at higher con-
centrations in t he stem. For all samples e xcept the specie
Pennisetum purpureum, the concentration of p-coumaric
acid is greater than that of ferulic acid, where the differ-
ence in concentration between the two acids is higher in
the stem t han in the leaf.
4. Conclusions
This study has demonstrated the potential use of an ul-
trasound bath for 120 minutes in the extr actio n of p heno-
lic acids in tropical forage. This method was fast and
reproducible when compared with the extraction tech-
nique using a t hermost atic bath with a programmed tem-
perature of 20˚C for 24 hours, which is the main method
repor ted in the literature for treatment of forage samples.
The samples were quantified using an internal standard
(o-coumaric acid). This method presented smaller values
of LOD, LOQ and RSD for the external standard.
5. Acknowledgements
The authors would like to thank the FAPEMIG (Fun-
dação de Amparo à Pesquisa do Estado de Minas
Gerais), CNPq (Conselho Nacional de Desenvolvimento
Cien- tífico e Tecnológico), CAPES (Coordenacão de
Aper- feiçoamento de Pessoal de Nível Superior) and
PRO- PESQ/UFJF (Pró-Reitoria de Pesquisa da
Universi- dade Federal de Juiz de Fora) for financial
support and grants Almeida, M. V. from the
Universidade Federal de Juiz de Fora for some
6. Referen ce
[1] R. Hatfield and R. S. Fukushima, “Can Lignin be Accu-
rately Measured?” Crop Science, Vol. 45, No. 3, 2005,
pp. 832-839. doi:10.2135/cropsci2004.0238
[2] C. D. Stalikas, “Extraction, Separation, and Detection
Methods for Phenolic Acids and Flavonoids,” Journal of
Separation Science, Vol. 30, No. 18, 2007, pp. 3268-3295.
[3] H. G. Jung, “Forage Lignins and their Effects on Fib er
Digestibility,” Agronomy Journal, Vol. 81, No. 1, 1989,
pp. 33-38.
[4] R. C. Sun, X. F. Sun and S. H. Zhang, “Quantitative De-
termination of Hydroxycinnamic Acids in Wheat, Rice,
Rye, and Barley Straws, Maize Stems, Oil Palm Frond
Fiber, and Fast-Growing Poplar Wood,” Journal of Agri-
cultural and Food Chemistry, Vol. 49, No. 11, 2001, pp.
5122-5129. doi:10.1021/jf010500r
[5] C. J. F. A. Brito, A. R. Rodella and F. C. Deschamps,
“Perfil Químico da Parede Celular e Suas Implicações na
Digestibilidade da Brachiara Brizantha e Brachiar a hu-
midicola,” Revista Brasileira de Zootecnia, Vol. 32, No.
6, 2003, pp. 1835-1844.
[6] M. D. Casler and H. G. Jung, “Relationships of Fibre,
Lignin, and Phenolics to in Vitro Fibre Digestibility in
Three Perennial Grasses,” Animal Feed Science Tech-
nology, Vol. 12 5, 20 06, pp. 151-161.
[7] F. C. Deschamps and L. P. Ramos, “Método para a
Determinação de Ácidos Fenólicos na Parede Celular de
Forragens,” Revista Brasileira de Zootecnia, Vo l . 31, No.
4, 2002, pp. 1634-1639.
[8] R. D. P. Hartley, “P-Coumaric and Ferulic Acid Compo-
nents of Cell Wall of Rygras s and thei r Relationship with
Lignin and Digestibility,” Journal of the Science of Food
and Agriculture, Vol. 23, No. 11, 1972, pp . 1 34 7-1354.
[9] H. G. Jung, “Maize Stem Tissues: Ferulate Deposition in
Developing Internode Cell Walls,” Phytochemistry, Vol.
63, No. 5, 2003, pp. 543-549.
[10] H. G. Jung and D. A. Deetz, “Cell Wall Lignification and
Degradability”. In: H.G. Jung, et al. (Eds.), Forage Cell
Wall Structure and Digestibility, AS A-CSSA-SSSA,
Madison, USA, 1993, pp. 315-346.
[11] J. R. Robbins, “Phenolic Acids in Foods: An Overvie w of
Analytical Methodology,” Journal of Agricultural and
Food Ch emistry, Vol. 51, N o. 10 , 2003, pp. 2866-2887.
[12] A. Carrasco-Pancorbo, A. M. Gómez-Caravaca, L. Cer-
retani, A. Bendini, A. Segura-Carretero and A. Fernán-
dez-Gutiérrez, “Rapid Quantification of the Phenolic
fraction of Spanish Virgin Olive Oils by Capillary Elec-
trophoresis with UV Detection,” Journal of Agricultural
and Food Chemistry, Vol. 54, No. 21, 2006, pp. 7984-
7991. doi:10.1021/jf0617925
[13] S. Ehala, M. Vaher and M. Kaljurand, “Characterization
of Phenolic Pro fi les of Northern European Berries b y Ca-
pillary Electrophoresis and Determination of their Anti-
oxidant Activity,” Journal of Agricultural and Food
Chemistry, Vol. 53, No. 16, 2005, pp. 6484-6490.
Copyright © 2011 SciRes. AJAC
[14] D. L. D. Lima, A. C. Duarte and V. I. Esteves, “Optimi-
zation of Phenolic Compounds Analysis by Capillary
Electrophoresis,” Talanta, Vol. 72, No. 4, 2007, pp.
1404-1409. doi:10.1016/j.talanta.2007.01.049
[15] Y. Y. Peng, J. N. Ye and J. L. Kong, “Determination of
Phenolic Compounds in Perilla Frutescens L. by Capil-
lary Electrophoresis with Electrochemical Detection,”
Journal of Agricultural and Food Chemistry, Vol. 53, No.
21, 2005, pp. 8141-8147. doi:10.1021/jf051360e
[16] A. Canini, D. Alesiani, G. D’Arcangelo and P. Tagliat-
esta, “Gas ChromatographyMass Spectrometry Analy-
sis of Phenolic Compounds from Carica Papaya L. Leaf,”
Journal of Food Composition and Analysis, Vol. 20, No.
7, 2007, pp. 584-590. doi:10.1021/jf051360e
[17] Y. C. Fiamegos, C. G. Nanos, J. Vervoort and C. D. Sta-
likas, “Analytical Procedure for the In-Vial Derivatiza-
tionExtraction of Phenolic Acids and Flavonoids in
Methanol ic and Aqueous Plant Extracts Followed by Gas
Chromatography with Mass-Selective Detection,” Jour-
nal of Chromatography A , Vol. 1041, No. 1 -2, 2004 , p p.
11-18. doi:10.1016/j.chroma.2004.04.041
[18] J. H. Grabber, J. Ralph and R. D. Hatfield,
“Cross-Linking of Maize Walls by Ferulate Dimerization
and Incorporation into Lignin,” Journal of Agricultural
and Food Chemistry, Vol. 48, No. 12, 2000, pp.
[19] M. Plessi, D. Bertelli and F. Miglietta, “Extraction and
Identification by GC-MS of Phenolic Acids in Traditional
Balsamic Vinegar from Modena,” Journal of Food Com-
position and Analysis, Vol. 19, No. 1, 2006, pp. 49-54.
[20] G. Sarath, L. M. Baird, K. P. Vogel and R. B. Mitchell,
“Internode Structure and Cell Wall Composition in Ma-
turing Tillers of Switchgrass (Panicum Virgatum. L),”
Bioresource Technology, Vol. 98, No. 16, 2007, pp.
2985-2992. doi:10.1016/j.biortech.2006.10.020
[21] Z. Spacil, L. Novakova and P. Solich, “Analysis of Phe-
nolic Compounds by High Performance Liquid Chroma-
tography and Ultra Performance Liquid Chromatogra-
phy,” Talanta, Vol. 76, 2008, pp. 189 -199.
[22] K. Chitindingu, A. R. Ndhlala, C. Chapano, M. A. Ben-
hura and M. Muchuweti, “Phenolic Compound Content,
Profiles and Antioxidant Activities of Amaranthus Hy-
bridus (Pigweed), B rachiaria Briz antha ( Upright Brachia-
ria) and Panicum Maximum (Guinea Grass),” Journal of
Food Biochemistry, Vol. 31, No. 2, 2007, pp.
206-216. doi:10.1111/j.1745-4514.2007.00108.x
[23] S. Gómez-Alonso, E. García-Romero and I. Hermosín-
Gutiérrez, “HPLC Analysis of Diverse Grape and Wine
Phenolics using Direct Injection and Multidetection by
DAD and Fluorescence,” Journal of Food Composition
and Analysis, Vol. 20, No. 7, 2007, pp. 618-626.
[24] Z. L. Huang, B. W. Wang, D. H. Eaves, J. M. Shikany
and R. D. P ace, “Phenolic Compound Pro file o f Selected
Vegetables Frequently Consumed by African Americans
in the Southeast United States,” Food Chemistry, Vol.
103, No. 4, 2007, pp. 1395-1402.
[25] C. Mertz, A. Gancel, Z. Gunata, P. Alter, C. Dhuique-
Mayer, F. Vaillant, A. M. Perez, J. Ruales and P. Brat,
“Phenolic Compounds, Carotenoids and Antioxidant Ac-
tivity of Three Tropical Fruits,” Journal of Food Compo-
sition and Analysis, Vol. 22, No . 5, 2009, pp. 381-387.
[26] M. A. M. Rodrigues, C. M. Guedes, J. W. Cone, A. H.
van Gelder, L. M. M. Ferreira and C. A. Sequeira, “Ef-
fec ts o f Phenolic Acid Structures on Meadow Hay Diges-
tibility,” Animal Feed Science Technology, Vol. 136,
2007, pp . 297-311.
[27] . C. I. G. Tuberoso, A. Kowalczyk, E. Sarritzu and P.
Cabras, “Determination of Antioxidant Compounds and
Antioxidant Activity in Commercial Oilseeds for Food
Use,” Food Chemistry, Vol. 103, No. 4, 2007, pp. 1494-
1501. doi:10.1016/j.foodchem.2006.08.014
[28] R. Al-Merey, M. S. Al-Masri and R. Bozou, “Cold Ultra-
sonic Acid Extraction of Copper, Lead and Zinc from
Soil Samples,” Analytica Chimica Acta, Vol. 452, No. 1,
2002, pp . 143-148. doi:10.1016/S0003-2670(01)01431-3
[29] K. Ashley, R. N. Andrews, L. Cavazos and M. Demange,
“Ultrasonic Extraction as a Sample Preparation Tech-
nique for Elemental Analysis by Atomic Spectrometry,”
Journal of Analytical Atomic Spectrometry, Vol. 16,
2001, pp . 1147-1153. doi:10.1039/b102027g
[30] A. Elik, “Ultrasound assisted pseudo-digestion of street
dust samples prior to determination by atomic absorption
spectrometry,” Talanta, Vol. 66, No. 4, 200 5, pp. 882-888.
[31] A. Marin, C. Lopez-Gonzales and C. Barbas, “Develop-
ment and Validation of Extraction Methods for Determi-
nation of Zinc and Arsenic Speciation in Soils Using Fo-
cused UltrasoundApplication to Heavy Metal Study in
Mud and Soils,” Analytica Chimica Acta, Vol. 4 42, 20 01 ,
pp. 305-318.
[32] S. C. C. Arruda, A. P. M. Rodriguez and M. A. Z. Arru-
da, “Ultrasound-Assisted Extraction of Ca, K and Mg
from in Vitro Citrus Culture,” Journal of the Brazilian
Chemical Societ y, Vol. 14, No. 3, 2003, pp. 470-474.
[33] M. Liva, R. Muñoz-Olivas and C. Câmara, “Determina-
tion of Cd in Sonicate Slurries and Leachates of Biologi-
cal and Environmental Materials by FI-CV-AAS,” Ta-
lanta, V ol. 51, No. 2, 2000, pp. 3 81-387.
[34] C. C. Nascentes, M. Korn and M. A. S. Arruda, “A Fast
Ultrasound-Assisted Extraction of Ca, Mg, Mn and Zn
from Vegetables,” Microchemical Jou rnal, Vol. 69, No. 1,
2001, pp . 37-43. doi:10.1016/S0026-265X(00)00192-2
[35] J. C. Cypriano, M. A. C. Matos and R. C. Matos, “Ultra-
sound-Assisted Treatment of Palm Oil Samples for the
Determination of Copper and Lead by Stripping Chrono-
potentiometry,” Microchemical Journal, Vol. 90, No. 1,
Copyright © 2011 SciRes. AJAC
2008, pp . 26-30. doi:10.1016/j.microc.2008.03.001
[36] E. A. Zakharova, V. I. Deryabina and G. B. Slepchenko,
“Optimization of the Voltammetric Determination of Ar-
senic in Foodstuffs,” Jounal of Analytical Chemistry,
Vol. 60, No. 6, 2005, pp. 503-507.
[37] P. J. Van Soest, “Nutritional Ecology of the Ruminant”,
Cornell University Press, New York, 1994.
Abbreviations Used
CE, capillary electrophoresis; GC-MS, gas chromatogra-
phy with a mass spectrometry detector; UPLC, ultra per-
formance liquid chromatography; HPLC, high perfor-
mance liquid chromatography; p-CA, ester-linked
p-coumaric acid; FA, ester-linked ferulic acid; LOD,
limit of detection; LOQ, limit of quantification; RSD,
relative standard deviation; CI, confidence interval.