Mass Spectrometry Reviews
REVIEW ARTICLE
Mass Spectrometry‐Based Applications in Tannin
Analytics: From Qualitative and Quantitative Analyses to
Biological Activity
Marica T. Engström1 | Maarit Karonen2
1Bioanalytical Laboratory, Institute of Biomedicine, University of Turku, Turku, Finland | 2Natural Chemistry Research Group, Department of Chemistry,
University of Turku, Turku, Finland
Correspondence: Marica T. Engström (mtengs@utu.fi)
Received: 8 January 2025 | Revised: 30 October 2025 | Accepted: 8 November 2025
Keywords: bioactivity | electrospray ionization | fragmentation patterns | high‐resolution mass spectrometry | multiple reaction monitoring
ABSTRACT
Tannins are widespread specialized plant metabolites that contribute significantly to the polyphenol content of plant‐based
diets. Their effects on human and animal health vary depending on their structure, with potential benefits including anti-
oxidative, antimicrobial, anthelmintic, and anticarcinogenic properties. Understanding tannin composition and quantity in
plant products is essential, as their bioactivities are influenced by their functional groups. Mass spectrometry‐based techniques
excel in tannin analysis, offering both qualitative and quantitative insights. Combining ultrahigh‐performance liquid chro-
matography with electrospray ionization and high‐resolution and triple quadrupole mass analyzers is optimal for compre-
hensive tannin profiling. Such an approach enables precise analysis and helps predict tannin bioactivities. This review
highlights the mass spectrometric analysis of proanthocyanidins and hydrolysable tannins, addressing ionization techniques,
interpretation of multiply charged ions, characteristic fragmentations, and reaction monitoring. Applications related to tannin
bioactivities are also briefly discussed, demonstrating the utility of mass spectrometry in tannin analysis in complex sample
matrices.
1 | Introduction
Tannins are a widely distributed heterogeneous group of plant
specialized metabolites. They occur ubiquitously in the plant
kingdom, for example, in bark, phloem, wood, stems, leaves,
seeds, fruits, fruit pods, berries, and nuts. Tannins can precipitate
proteins, and historically, they are among the first natural
products utilized industrially, that is, as tanning agents for
transformation of fresh hides into leather (L. J. Porter 1989a; L. J.
Porter 1989b). Nowadays, plant tannins are known to have many
beneficial bioactivities, for example, antioxidant, anti-
carcinogenic, anthelmintic, and antimicrobial activities
(Engström et al. 2019; Engström, Karonen et al. 2016; Gaudin
et al. 2016; Gong et al. 2018; Gontijo et al. 2019; Hoste et al. 2006;
Karonen et al. 2020; Kolodziej et al. 1999; López‐Andrés
et al. 2013; Puljula et al. 2020; Quijada et al. 2015; Reddy
et al. 2007; Scalbert 1991; Soorkia et al. 2020). Tannins can also
be modified or transformed, for example, by alkaline or aerobic
oxidation (García et al. 2016; Imran et al. 2021; Karonen
et al. 2021; Petit et al. 2013), by synthetic approaches
This is an open access article under the terms of the Creative Commons Attribution‐NonCommercial License, which permits use, distribution and reproduction in any medium, provided the original
work is properly cited and is not used for commercial purposes.
© 2025 The Author(s). Mass Spectrometry Reviews published by Wiley Periodicals LLC.
Abbreviations: APCI, atmospheric pressure chemical ionization; APPI, atmospheric pressure photo ionization; CI, chemical ionization; DART, direct analysis in real time; DESI, desorption
electrospray ionization; EI, electron ionization; ESI, electrospray ionization; FT‐ICR, Fourier transform ion cyclotron resonance mass analyzer; HPLC, high‐performance liquid chromatography;
HT, hydrolysable tannin; LC, liquid chromatography; MALDI, matrix assisted laser desorption/ionization; MRM, multiple reaction monitoring; MS, mass spectrometry; MS/MS, tandem mass
spectrometry; PA, proanthocyanidin; Q, quadrupole; QqQ, triple quadrupole; SRM, single reaction monitoring; TOF, time‐of‐flight mass analyzer; UHPLC, ultrahigh‐performance liquid
chromatography.
1Mass Spectrometry Reviews, 2025; 1–27
https://doi.org/10.1002/mas.70013
(Cobo et al. 2024; Deffieux et al. 2011; Laitila et al. 2023; Malik
et al. 2012; Petit et al. 2019; Pouységu et al. 2011; Quideau
et al. 2011; Sylla et al. 2015) or during wine aging (He et al. 2010;
Jourdes et al. 2011; Laitila and Salminen 2020; Quideau
et al. 2003; Rasines‐Perea et al. 2019; Saucier et al. 2006).
The tannin composition of a single plant species can be complex
(Salminen and Karonen 2011). Even the organs of the same
plant species may produce very variable tannin structures
(Tuominen et al. 2013). On the basis of the major differences in
their structures, terrestrial tannins are divided into two groups:
proanthocyanidins (PAs, syn. condensed tannins) and hydro-
lysable tannins (HTs). PAs are oligomers (two to ten monomeric
units) and polymers (> 10 monomeric units) of flavan‐3‐ol units
which have variation in their hydroxylation patterns, stereo-
chemistry at C2 and C3, degree of polymerization, sequential
order of the flavan‐3‐ol units and location and stereochemistry
of interflavanoid bonds (Figure 1, Hemingway 1989a). Mono-
meric units can be linked by C4→ C8 and/or C4→ C6 (B‐type
PAs) or doubly linked with an additional C2→O→C7 ether
bond (A‐type PAs). The most common PAs are procyanidins
containing (+)‐catechin with 2R,3S stereochemistry and (‐)‐
epicatechin with 2R,3R stereochemistry as monomeric units
and prodelphinidins containing (+)‐gallocatechin and (‐)‐
epigallocatechin. In addition to the six main classes, some less
common PA classes exist, such as proapigenidins and prolute-
olinidins (Hemingway 1989a; Krueger et al. 2003; L. J.
Porter 1989a; L. J. Porter 1989b). In plants, individual PA
structures often contain various types of subunits; for instance,
pure procyanidin or prodelphinidin oligomers or polymers are
less common than their mixtures.
HTs are typically divided into three major subclasses, that is,
simple gallic acid derivatives, gallotannins and ellagitannins.
Simple gallic acid derivatives most commonly have either glu-
cose or quinic acid as the central polyol galloylated with
monogalloyl groups (Figure 2A), whereas gallotannins also
contain digalloyl or trigalloyl groups in their structures. Ella-
gitannins can be further classified into multiple subgroups,
these seven being the most common groups for ellagitannin
monomers: hexahydroxydiphenoyl esters (Figure 2B),
dehydrohexahydroxydiphenoyl esters and their modifications
(Figure 2C,D), gallagyl esters (Figure 2E), non-
ahydroxytriphenoyl esters (Figure 2F), and flavanoellagitannins
(Figure 2G). The diversity of ellagitannin structures is multi-
plied by their various types of oligomerization, a feature typi-
cally not found with simple gallic acid derivatives or
gallotannins.
2 | Different Ionization Techniques for Tannin
Analysis
In the mass spectrometric analysis of tannins, two important
choices must be made. The first one is related to the ionization,
that is, the selection of the ionization technique used, and
the second is related to the actual mass analysis, that is, the
selection of mass analyzer used. There are several suitable
ionization techniques for tannins. The early ionization tech-
niques, such as electron ionization (EI), are poorly suited for
tannin analysis (Cheynier and Fulcrand 2003). EI requires
volatile analytes, but tannins are thermosensitive and non-
volatile in their native form. However, this limitation can be
overcome by derivatization, for example by using methylation
by dimethyl sulfite (Nonaka et al. 1983). In addition, EI is a
hard ionization technique and with typical ionization energy of
70 eV induces extensive fragmentation in which case the
detection of molecular ions can be challenging. Later ionization
techniques, such as fast atom bombardment and liquid sec-
ondary ion mass spectrometry, which generate the secondary
ions when the sample is irradiated with energetic neutral atoms
in fast atom bombardment or primary beam of ions in liquid
secondary ion mass spectrometry, have been mainly used for
the analysis of purified tannins as the sample must be dissolved
in a matrix. For example, PAs have been analyzed using glyc-
erol (Freitas et al. 1998), thioglycerol (Vivas et al. 1996), or
eutectic mixture of dithiothreitol and dithioerythritol
(Barofsky 1989; Karchesy et al. 1986) as matrices.
Matrix also plays an important role in matrix‐assisted laser
desorption‐ionization (MALDI) of tannins. The use of MALDI
techniques involves the optimization of many factors, such as
FIGURE 1 | Model structure and numbering of a B‐type proanthocyanidin with C4→ C8 linkages.
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the selection of an appropriate matrix, optimal sample prepa-
ration including the mixing and drying or crystallization of
matrix and sample, the actual ionization parameters related, for
example, to the laser power and behavior of the matrix during
laser irradiation, and the selection of calibration standards
(Behrens et al. 2003; Es‐Safi et al. 2006; Reed et al. 2005). The
advantages of MALDI are its sensitivity, a single ionization
event and the infinitesimal fragmentation of tannins when
FIGURE 2 | Model structures of monomeric hydrolysable tannins: (A) pentagalloylglucose, (B) tellimagrandin I, (C) geraniin, (D) chebulagic
acid, (E) punicalagin, (F) vescalagin and (G) stenophyllanin A.
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main target is the determination of molecular formula (Krueger
et al. 2000; Yang and Chien 2000). If controlled fragmentation is
desired, this can be obtained by using postsource decay frag-
mentation, which allows the detailed characterization and
sequencing of individual PA chains, that is, a sequential loss of
one monomeric unit after the other (Behrens et al. 2003). This
technique allows the selection of a precursor ion, for example
an [M+Na]+ ion for a procyanidin tetramer at m/z 1177, in a
distinct mass window and the subsequent analysis of its frag-
ment ions, in the case of procyanidin tetramer signals at m/z
889 and 887 and at m/z 601 and 599 corresponding to trimeric
and dimeric fragments, as presented in Figure 3, respectively
(Behrens et al. 2003). MALDI techniques have their own ben-
efits and challenges related to direct tannin analyses as previ-
ously reviewed by Flamini (2003), Reed et al. (2005) and
Monagas et al. (2010).
Atmospheric pressure ionization techniques, such as atmo-
spheric pressure chemical ionization (APCI), atmospheric
pressure photo ionization (APPI), and electrospray ionization
(ESI), can be easily combined with liquid chromatographic
techniques. APCI is rarely used for tannins as it is most suitable
for thermally stable compounds with molecular weights less
than 1500 Da as it produces mainly singly charged ions (de
Hoffmann and Stroobant 2001; Holčapek et al. 2010). APCI has
been used, for example, for the analysis of flavan‐3‐ols, PA
dimers, trimers and their gallates, modified tannins in wines
and dimeric and trimeric ellagitannins (de Pascual‐Teresa
et al. 2000; Franceschi et al. 2011; Maillard et al. 1996;
Mazerolles et al. 2010; Weber et al. 2007). Based on literature
search, APPI is even less used and to our knowledge it has been
applied only for tannins in soil samples to complement ESI
(Vinci et al. 2022). However, based on our experience ESI is a
superior ionization technique regarding tannin analyses. Its
ability to produce multiply charged ions folds up the m/z scale
by the number of charges, shifting the ions into an m/z range
below 3000, which is accessible by most mass analyzers
(Gross 2017). In ESI, the analyte solution is supplied through a
thin spray capillary, nebulizer needle, which is kept at high
potential of 3–6 kV (de Hoffmann and Stroobant 2001) and
sheathed with nebulizer gas to form an electrically charged
aerosol. When the analyte solution comes out from the capil-
lary, it breaks up to form highly charged droplets. The ions
present in the charged droplets are liberated into gas phase, via
several evaporation (droplet shrinking) and droplet dis-
integration steps, either according to charge‐residue model or
ion evaporation model (Gross 2017), yielding multiply charged
ions for larger molecules. In general, ESI is a soft, rapid, sen-
sitive technique which can be used either by direct infusion, as
nicely outlined previously (Hayasaka et al. 2003; Roux
et al. 1998), or in combination with HPLC or UHPLC tech-
niques (Baert et al. 2015; Hammerstone et al. 1999; Karonen
et al. 2011, 2010, 2004; Lazarus et al. 1999; Rauha et al. 2001;
Salminen et al. 1999). Multiply charged ions obtained by ESI‐
MS enables the detection and characterization of high‐
molecular weight tannins, including PAs with a degree of
polymerization of up to 22 (Es‐Safi et al. 2006; Karonen
et al. 2011) and undecameric ellagitannins with a molecular
weight of 8624 Da (Salminen et al. 2011). Although ESI is a soft
ionization technique, it produces some fragment ions for tan-
nins. Therefore, the different monomeric units in oligomeric
tannins can be detected, see for example Karonen et al. (2010)
for HT oligomers, and Karonen et al. (2011) for PA oligomers.
For PAs, ESI‐MS allows the determination of the nature and
FIGURE 3 | Postsource decay matrix assisted laser desorption/ionization—time‐of‐flight mass spectrum of a procyanidin tetramer from lime
(Tilia cordata) together with the possible fragmentation pattern in positive ion mode. The colors of the brackets refer to the corresponding
fragmentation site drawn with a dashed line of the same color. The mass spectrum is reprinted from Behrens et al. (2003) with permission from
Elsevier. [Color figure can be viewed at wileyonlinelibrary.com]
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proportions of constitutive units, degree of polymerization, type
of the interflavanoid linkages (A‐ or B‐type), and the nature and
proportion of substituents as discussed by Cheynier and
Fulcrand (2003). A‐type PAs can be recognized by their
molecular ions corresponding to the loss of two hydrogens
during the biosynthetic formation of the additional ether bond
in comparison with B‐type PAs. Therefore, the m/z values of
A‐type PAs differ by 2 Da from the corresponding B‐type ones.
The substituents in PA structures identified by ESI can be, for
example, xylose (Liimatainen et al. 2012), glucose (Salminen
et al. 2005), or galloyl group (Karonen et al. 2021). In general,
tannins are better ionized in the negative mode in comparison
to positive mode due to the acidity of phenolic hydroxyl groups.
For example, PA pentamers seem to be the limit of PA detection
when positive ESI is used (Hammerstone et al. 1999). We have
also experimentally observed that the use of positive ESI pro-
duces more in‐source fragment ions in comparison to negative
ESI which might hinder the correct identification of the
molecular ion. The negative ionization gives also simpler mass
spectra due to the absence of intensive adduct ions (de
Hoffmann and Stroobant 2001; Hayasaka et al. 2003). Typical
adducts in positive ion mode are formed with metal ions (e.g.,
Na⁺, K⁺), solvent molecules (e.g., H₂O), and counter ions (e.g.,
Cl⁻, AcO⁻) which can complicate mass spectra by generating
additional peaks.
Desorption electrospray ionization (DESI) introduced in 2004
and direct analysis in real time (DART) introduced in 2005 have
been less used so far, but their applications for rapid visual-
ization or imaging of monomers and small oligomers seem
promising. For example, tannins among other metabolites have
been imaged via imprint, followed by DESI/postphotoionization
(imprint DESI/PI) mass spectrometry imaging (Wu et al. 2022).
The technique also allows the comprehensive mapping of the
metabolomic network of catechin in fresh tea leaves. DESI has
also been used for rapid visualized characterization of tannins
together with other phenolics in tea extracts after their sepa-
ration by high‐performance thin‐layer chromatography (Tang
et al. 2021). DART is a soft ionization technique suitable for the
fast detection of flavan‐3‐ols. For example, eucalyptus leaf and
stem tissues were placed directly in the gap between the
skimmer of DART ion source and the capillary inlet of time‐of‐
flight (TOF) mass analyzer which enabled the ionization and
analysis of catechin among other compounds (Maleknia
et al. 2009). Similarly, epiafzelechin, the monomeric building
block of propelargonidin, was successfully ionized from a root
sample of Cassia sieberiana placed in the open ion source
(Kpegba et al. 2011). Ambient desorption/ionization methods
are superior in many ways as they are directly applicable to
solids, liquids, frozen solutions or complex biological materials,
but they have a limited potential for quantification as the
detection of the compound heavily depends on the matrix
(Gross 2017).
3 | Importance of High‐Resolution in Qualitative
Tannin Analysis
Different mass analyzers have different roles in tannin analysis.
Due to their low‐resolution, quadrupole instruments are not
suitable for accurate mass analysis but give superior qualitative
and quantitative results when tandem mass spectrometric
methods, such as multiple reaction monitoring, are applied.
These applications are discussed in detail in Section 6, here the
focus is on high‐resolution mass analyzers that excel at tannin
characterization due to their ability to provide detailed isotopic
patterns (Gross 2017). Most commonly orthogonal‐acceleration
TOF or orbitrap mass analyzers are used as they can be con-
veniently combined with LC, see for example, Karonen et al.
(2011), Lin et al. (2014), Navarro‐Hoyos et al. (2017) and Leppä
et al. (2018). The main advantages of TOF instruments for
tannins are the same as for all TOF analysis: the instrument
construction is straightforward and allows accurate mass mea-
surements, the m/z range of a TOF analyzer is in theory
unlimited; in addition, TOF analyzers have high sensitivity
through high ion transmission and the mass spectrum acqui-
sition rate of TOF is very high (Gross 2017). Similarly, the ad-
vantages of using an orbitrap for tannin analyses are the same
as for other compounds: ultrahigh‐resolution, high mass accu-
racy, and increased speed and sensitivity, especially when
combined with a bent RF‐only quadrupole C‐trap (Gross 2017).
Typically, the resolving power of TOF and orbitrap varies
widely based on their design, mode of operation and intended
use. Generally, TOF can achieve the resolving power to
R > 30,000 and orbitrap the resolving power of
R = 20,000–200,000 (Gross 2017). When a TOF or an orbitrap is
combined with a quadrupole, tandem MS experiments can be
used with high‐resolution and mass accuracy. In addition, a few
studies have utilized Fourier transformation ion cyclotron re-
sonance (FT‐ICR) mass spectrometers for tannins or tannin‐like
compounds (Cooper and Marshall 2001; Ferreira et al. 2014; He
et al. 2019). The operating principle of FT‐ICR is similar to
orbitrap as they both employ image current detection of ion
oscillations and Fourier transformation for the conversion of
the transient into the frequency domain. However FT‐ICR
operates with a magnetic field, resulting in higher costs and
more stringent requirements (installation room, power con-
sumption, air conditioning load, personnel qualification, and
maintenance) compared to orbitrap which utilizes an electro-
static field (Gross 2017).
The isotopic patterns obtained by high‐resolution mass spectro-
meters enable the accurate mass calculations for each singly or
multiply charged ion and thereby provides the exact mass of the
corresponding tannin. The charge state of the ion is determined by
the mass difference of adjacent isotopic peaks, and the multiply
charged ions and their charge states can be easily distinguished
from the singly charged ones. Tannins contain H, C and O which
are all isotopic elements, but only the abundance of 13C (1.08%) is
significant from the isotopic pattern point of view. The isotopes 2H
(0.01%) and 17O (0.04%) and 18O (0.21%) have so low abundances
that they do not affect the isotopic pattern significantly. In the
mass spectrum, the information of the analyte ions is displayed at
the m/z scale, and the mass difference between the isotopic peaks
can be described as Δ (m/z) = 1/z and thereby, the charge state
calculated based on the observed mass difference as z=1/Δ (m/z).
As an example, two oligomeric ellagitannins both with molecular
ions at m/z 934.07 (sanguiin H‐6 and lambertianin C) are shown
in Figure 4A,B. When the isotopic distributions of the ions are
zoomed, different isotopic patterns can be detected (Figure 4C,D).
By accurate mass data, the charge states of the ions are first
determined: for sanguiin H‐6 (Figure 4C), Δ (m/z) is 0.5 = 1/2
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corresponding to z=2, i.e., the doubly charged [M–2H]2– ion, and
for lambertianin C (Figure 4D), Δ (m/z) is 0.33 = 1/3 corre-
sponding to z=3, i.e., the triply charged [M–3H]3– ion. Then, the
corresponding exact masses and molecular formulae based on the
monoisotopic peak are calculated.
As the size of the tannin (and the number of carbon atoms)
increases, the probability that an ion contains at least one 13C
becomes larger. Therefore, the monoisotopic peak is smaller for
the trimeric lambertianin C (C123H80O78; Figure 4D) in com-
parison to the one of dimeric sanguiin H‐6 (C82H54O52;
Figure 4C). When the molecular size is large enough, as illus-
trated in Figure 5 with an example of Lysimachia vulgaris PAs
exhibiting the degree of polymerization of 17 (Leppä
et al. 2018), it might be challenging to detect the monoisotopic
peak. In such a case, it is helpful to use the simulation tools of
mass spectrometric software to simulate the isotopic distribu-
tion and compare it with the measured one. In the example
shown, the experimental data contain many isotopic PA pat-
terns in addition to background ions (Figure 5A). However,
with the isotope simulation the correct monoisotopic peaks can
be delineated and found to consist of 3 procyanidin and 14
prodelphinidin units (Figure 5B) and of 4 procyanidin and 13
prodelphinidin units (Figure 5C).
It must also be noted that polyphenols can be oxidized into
quinone forms during the mass spectrometric analysis and,
therefore, there can be minor signals in the mass spectra of
polyphenols having the m/z values 2 Da smaller than the cor-
responding polyphenol (Karonen et al. 2021). However, these
minor peaks caused by the oxidation of tannins should not be
mixed with A‐type PAs having an additional ether bond and a
mass difference of 2 Da in comparison to B‐type PAs. Figure 6A
shows an example of procyanidin dimer having only one B‐type
linkage (578 Da): the mass spectrum exhibits the deprotonated
molecule at m/z 577 and the minor signal corresponding to the
oxidation product formed during the analysis at m/z 575. In
Figure 6B, the corresponding procyanidin dimer with both A‐
and B‐type linkages (576 Da), exhibiting a deprotonated mole-
cule at m/z 575, can be seen along with minor signals corre-
sponding to the oxidation product at m/z 573.
Despite these few things to be considered in the interpretation of
mass spectra, it is still important to remember that multiply
charged ions are an advantage in the tannin analysis, as they
significantly expand the available mass range. For example, a
previous work showed detection of procyanidin polymers up to
the degree of polymerization of 22 by using ESI‐QTOF‐MS: the
singly charged ions were dominant for dimers and trimers (578
and 866 Da), doubly charged ions for tetra‐ to octamers
(1154–2306Da), triply charged ions for nona‐ to octadecamers
(2594–5187Da) and fourfold charged ions were detected from the
hexadecamers onwards (> 4611Da, (Karonen et al. 2011). When
the multiply charged ions are analyzed with ultrahigh‐resolution,
very complex patterns can be accurately characterized for oligo‐
and polymeric PAs isolated from Lysimachia vulgaris extract as
FIGURE 4 | Molecular ions of (A) sanguiin H‐6 and (B) lambertianin C with their isotopic distributions (C and D), respectively, highlighting the
mass differences between the isotopic peaks in negative ion mode ESI‐MS. When the data is examined with a precision of two decimal places (A and
B), the mass spectra looks the same, but when the isotopic distributions are zoomed and more precise numerical values are viewed (C and D), a clear
difference between the mass spectra is noticed. [Color figure can be viewed at wileyonlinelibrary.com]
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shown in Figure 7 (Leppä et al. 2018). The sample contained a
diverse mixture of B‐type PAs consisting of both procyanidin and
prodelphinidin units. Procyanidin oligomers can be observed at
m/z 577.13 + 288.06 × n (where n is the number of additional
monomeric units attached to the dimer) when they are present as
singly charged ions, at m/z 576.13+ 144.03 × n when doubly
charged and at m/z 959.87 + 96.02 × n when triply charged and,
similarly, prodelphinidin oligomers at m/z 609.12 + 304.06 × n
when singly charged, at m/z 608.12+ 152.03 × n when doubly
charged and at m/z 1013.19 + 101.35 × n when triply charged.
The same systematic approach can be applied when a
procyanidin unit is replaced with a prodelphinidin unit (having
an additional hydroxyl group) in the PA oligomer. In this case,
there is an increment in the m/z value of the ion depending on
the charge state of the ions: this increment is +16.00 Da for singly
charged ions, +8.00 Da for doubly charged ions, +5.33 Da for
triply charged ions and +4.00Da for fourfold charged ions
(Leppä et al. 2018).
The exact masses are needed for correct structural identification
and molecular formulae. The differences between the molecular
weights of tannins can be minor although the tannin structures
are totally different. This phenomenon can be illustrated by
comparing a galloylated dimeric PA structure consisting of two
procyanidin units and two galloyl groups (molecular formula
C44H34O20, Mcalculated 882.16435) with a trimeric PA consisting
FIGURE 5 | (A) Experimental ESI‐MS data in negative ion mode of heptadecameric proanthocyanidins of Lysimachia vulgaris with the simulated
isotopic patterns of heptadecamers consisting of (B) 3 procyanidin and 14 prodelphinidin units and (C) 4 procyanidin and 13 prodelphinidin units.
[Color figure can be viewed at wileyonlinelibrary.com]
FIGURE 6 | Mass spectra of procyanidin dimers having (A) only one B‐type linkage and (B) both A‐ and B‐type linkages obtained by negative ion
mode ESI‐MS.
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of two procyanidin and one prodelphinidin units (molecular
formula C45H38O19, Mcalculated 882.20074) (Karonen et al. 2021).
The dimer has a deprotonated molecule at m/z 881.15267 and
the trimer at m/z 881.19452 (Figure 8) and these m/z values
cannot be separated from each other by low‐resolution mass
analyzers. In addition, cluster and fragment ions can be
observed. The fragmentation of tannins is discussed in detail in
Chapter 4 (PAs) and Chapter 5 (HTs).
4 | Characteristic Fragmentations of
Proanthocyanidins
PAs have very characteristic and well‐known fragmentation
patterns in negative ion mode ESI‐MS analysis (Friedrich
et al. 2000; Gu et al. 2003a; Karonen et al. 2004). The frag-
mentation of heterocyclic ring occurs via retro‐Diels–Alder
fragmentation or heterocyclic ring fission (Figure 9). The retro‐
FIGURE 7 | High‐resolution mass data of Lysimachia vulgaris proanthocyanidins (Leppä et al. 2018) obtained by negative ion mode ESI‐MS. DP
refers to degree of polymerization, PC to procyanidin unit and PD to prodelphinidin unit. The isotopic patterns of the ions at m/z 865 and 1201 are
zoomed to highlight the detailed information available. *The ions are [M–2H]2– ions. **The ions are [M–3H]3– ions and the m/z values on the top of
the peaks refer to the most intensive isotopic peaks, not to the monoisotopic masses. [Color figure can be viewed at wileyonlinelibrary.com]
FIGURE 8 | Ultrahigh‐resolution mass spectra of (A) a galloylated procyanidin dimer and (B) a proanthocyanidin trimer consisting of two
procyanidin and one prodelphinidin units exhibiting almost similar molecular ions in negative ion mode ESI‐MS.
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Diels–Alder fragmentation has been considered to be the most
important fragmentation reaction as it allows the reliable
characterization of PA dimers. The retro‐Diels–Alder fragmen-
tation of the extension unit produces fragment ions with larger
π–π hyperconjugated systems than the fragmentation of the
terminal unit and is therefore considered to be energetically
more favorable (Friedrich et al. 2000; Gu et al. 2003a; Karonen
et al. 2004). Typically, the retro‐Diels–Alder fragmentation is
followed by subsequent elimination of water (−18 Da). Some-
times, the abundance of the water loss product is higher than
FIGURE 9 | Fragmentation pathways of (A) A‐type and (B) B‐type procyanidin dimers in negative ion mode ESI‐MS. The fragmentation
mechanisms are retro‐Diels–Alder fragmentation (RDA), heterocyclic ring fission (HRF) and quinone methide cleavage (QM). The unstable
intermediate ions are marked with brackets. Structures shown are neutral molecules; the m/z values below correspond to the detected ions. [Color
figure can be viewed at wileyonlinelibrary.com]
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that of the retro‐Diels–Alder product ion (Friedrich et al. 2000).
The interflavanoid bond cleaves via quinone methide cleavage
or direct cleavage which does not include charge or radical
driven rearrangement. The fragmentation patterns are similar
for both A‐type and B‐type PAs as presented for procyanidin
dimers in Figure 9 (Friedrich et al. 2000; Gu et al. 2003a;
Karonen et al. 2004; Rue et al. 2018; Sui et al. 2016). In retro‐
Diels–Alder fragmentation, the catechol unit is cleaved pro-
ducing the ions with m/z 423 (A‐type) and m/z 425 followed by
the water elimination producing the ion at m/z 407 (B‐type). In
heterocyclic ring fission, a phloroglucinol unit is cleaved
resulting in the ions with m/z 449 (A‐type) and m/z 451 (B‐
type). The quinone methide cleavage produces the ions with m/
z 285 (A‐type) and m/z 287 (B‐type) for the extension units and
the ion with m/z 289 for the terminal units.
The fragmentation patterns are similar for oligomers and
polymers but they tend to yield numerous smaller fragment
ions rather than a few larger ones (Gu et al. 2003b; Karonen
et al. 2004). This also makes the interpretation of their mass
spectra challenging as it may be difficult to know with certainty
whether an ion is an intact deprotonated molecule or whether it
is a fragment ion. Therefore, a good chromatographic separa-
tion can support the interpretation of MS data and characteri-
zation of PAs; for this purpose diol columns and hydrophilic
interaction chromatography are beneficial (Kalili and de
Villiers 2010, 2009; Karonen et al. 2011; Kelm et al. 2006). In
hydrophilic interaction chromatography‐ESI‐MS analysis, PAs
elute according to increasing degree of polymerization and the
sensitivity is enhanced in ESI due to the high proportion of
organic solvent used in the gradient (Kalili and de
Villiers 2010, 2009; Karonen et al. 2011). In reversed phase
liquid chromatographic analysis, the elution order is not
directly related to the degree of polymerization. The following
order has been generally observed for flavan‐3‐ols, procyanidin
dimers and procyanidin trimers (de Pascual‐Teresa et al. 1998;
Escribano‐Bailon et al. 1992; Zhao et al. 1999):
C2 B3 B1 catechin B6 B4 B2
epicatechin B7 C1 B5
< < < (+) − < < <
< (−) − < < < ,
where C2=catechin‐(4α→8)‐catechin‐(4α→8)‐catechin, B3=
catechin‐(4α→8)‐catechin, B1=epicatechin‐(4β→8)‐catechin,
B6=catechin‐(4α→6)‐catechin, B4=catechin‐(4α→8)‐epicatechin,
B2=epicatechin‐(4β→8)‐epicatechin, B7=epicatechin‐(4β→6)‐
catechin, C1=epicatechin‐(4β→8)‐epicatechin‐(4β→8)‐epicatechin,
and B5=epicatechin‐(4β→6)‐epicatechin.
Only shorter PA oligomers can be separated and defined as
sharp peaks by reversed phase liquid chromatography. Poly-
meric PAs are detected by the formation of characteristic large
unresolved “humps” in the chromatograms (Adamson
et al. 1999; Guyot et al. 1998, 1997; Roux et al. 1998). On the
other hand, the challenges of poor separation can be overcome
by single and multiple reaction monitoring (SRM and MRM,
respectively) methods utilizing a specific PA fragmentation as
discussed in Section 3. Even though PAs can be characterized
by mass spectrometric techniques, the position and stereo-
chemistry of the interflavanoid linkages (C4→ C8 or C4→ C6)
cannot be identified (Friedrich et al. 2000). Moreover, the
differentiation of stereoisomers needs additional information,
such as the retention order in LC‐MS analysis. For example,
monomeric flavan‐3‐ols are known to elute in the following
order in reversed phase liquid chromatographic analysis
(Koupai‐Abyazani et al. 1992):
gallocatechin epigallocatechin
catechin
epicatechin
(+) − < (−) −
< (+) −
< (−) − .
Monomeric units also have their own characteristic product
ions, that is, m/z 245 for procyanidin unit or m/z 261 for
prodelphinidin unit, resulting from the cleavage of the hetero-
cyclic ring by the loss of the –CH2–CHOH– group (Gariboldi
et al. 1998; Pérez‐Magariño et al. 1999).
5 | Characteristic Fragmentations of
Hydrolysable Tannins
The fragmentation patterns of HTs are well studied and docu-
mented. However, there are some variations in the reported
fragment ions, which mainly stem from the different instru-
mentation and analysis conditions used (Mena et al. 2012). The
fragmentation patterns of negatively charged galloylglucoses
and gallotanninss have been studied by utilizing full scan MS
(e.g., Barry et al. 2001; Mämmelä et al. 2000; Salminen
et al. 1999) but tandem mass spectrometry has enabled more
detailed studies on their fragmentation patterns. Galloylglu-
coses and gallotannins contain two types of acyl moieties that
can be differentiated upon fragmentation. The current knowl-
edge of fragmentation patterns of galloylglucoses is summarized
in Figure 10. The most characteristic fragment ion for galloyl-
glucoses is a gallate anion at m/z 169 which stems from the
fragmentation of the galloyl groups attached to the core polyol
by ester bonds as gallate ions (Figure 10). Alternatively, the
galloyl group may be eliminated as a neutral fragment (152 Da).
For example, pentagalloylglucose yields repetitive loss of galloyl
moieties (152 Da, Figure 10) resulting in tetra‐, tri‐, di‐, and
monogalloylglucose ions (Barry et al. 2001; Salminen et al. 1999;
Singh et al. 2016; Hooi Poay et al. 2011). Other fragmentations
include the loss of water (−18 Da) and decarboxylation
(−44 Da). For the smaller galloylglucoses, di‐ and mono-
galloylglucoses, the additional losses of 60 Da (Barry et al. 2001;
Soong and Barlow 2005; Zywicki et al. 2002) is caused by the
cross‐ring fragmentation of a glucose unit (Meyers et al. 2006;
Sobeh et al. 2019), for example via removal of formaldehyde
moieties (Hooi Poay et al. 2011; Taylor et al. 2005) as presented
in Figure 10.
In gallotannins, the depside bonds between the core galloyls
and additional galloyl groups are less stable than the ester
bonds between the core polyol and therefore undergo frag-
mentation before the core galloyl groups, leading to the loss
of 152 Da (Berardini et al. 2004; Luo et al. 2014; Regazzoni
et al. 2013; Salminen et al. 1999). After the removal of these
additional galloyl groups, the fragmentation of the remain-
ing galloylglucose structure follows the patterns presented
above.
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In comparison to galloylglucosess and gallotanninss, ellagi-
tannins are structurally more diverse and correspondingly, their
fragmentation patterns in negative ESI‐MS are more complex.
For primary hexahydroxydiphenoyl esters, such as tell-
imagrandin II (Figure 2B), the main fragment ions are often
related to the galloyl group and the hexahydroxydiphenoyl
group, yielding fragment ions of m/z 169 (Figure 10) and m/z
301 (Figure 11), respectively (Fracassetti et al. 2013; Moilanen
et al. 2013; Mullen et al. 2003; Regueiro et al. 2014; Zywicki
et al. 2002). For the further fragmentation, fragment ions at m/z
275,m/z 257,m/z 229 andm/z 145 have been reported (Aguilar‐
Zárate et al. 2017; Bowers et al. 2018; Del Bubba et al. 2012;
Duckstein et al. 2013; Engström et al. 2015; Mullen et al. 2003;
Singh et al. 2016; Ventura et al. 2024; Yang et al. 2012). Other
functional groups also experience characteristic fragmentations,
which is a great benefit for structural determination. For ex-
ample, punicalagin (Figure 2E) with an hexahydroxydiphenoyl
group and a gallagyl group, yields fragment ions at m/z 301
(hexahydroxydiphenoyl), m/z 601 (gallagyl) and m/z 781
(gallagyl‐glucose) as presented in Figure 12A (Mena et al. 2012;
Pfundstein et al. 2010; Seeram et al. 2005). For dehy-
droellagitannins that contain dehydrohexahydroxydiphenoyl
group, such as geraniin (Figure 2C), loss of water from the
deprotonated molecule at m/z 951 is characteristic, giving a
fragment ion at m/z 933 for geraniin (Palanisamy et al. 2011). If
the ET contains a chebuloyl group instead (Figure 2D), diag-
nostic fragment ion at m/z 337 [Chebulic acid–H‐H2O]– is
formed. In addition, fragment ions at m/z 319, m/z 293 and m/z
FIGURE 10 | The fragmentation pathways of galloylglucoses in negative ion mode ESI‐MS. GG, galloylglucose.
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275 are formed due to consequent losses of H2O or CO2 as
shown in Figure 12B (Pfundstein et al. 2010; Yang et al. 2012).
The loss of water is a characteristic fragmentation also for some
other ellagitannins and this feature can be used for example to
differentiate between the different orientations of the hydroxyl
group at the C1 position of the C‐glycosidic ellagitannin isomers
(e.g., vescalagin vs. castalagin, see Figure 2F). The loss of water
is seen for vescalagin type β‐OH (Figure 11) but not castalagin
type α‐OH C‐glycosidic ellagitannins due to steric and intra-
molecular stabilization effects (Moilanen et al. 2013; Quideau
et al. 2005, 2003; Yoshida et al. 1991). Regarding other frag-
mentations, the nonahydroxytriphenoyl group is rather stable,
and therefore vescalagin yields characteristic fragment ions at
m/z 631 (vescalin) and m/z 613 (vescalin‐H2O) (Figure 11). If
the carboxylic acid group is free in the ET structure, as it is, for
example, in vescavaloninic acid and castavaloninic acid, the
cleavage of –COOH (−44 Da) can be detected (Moilanen
et al. 2013). In addition, flavanoellagitannins have the charac-
teristic cleavage of the flavan‐3‐ol moiety (Figure 2G) exhibiting
a fragment ion at m/z 289.
For oligomeric ellagitannins, the individual monomeric units
can be detected (Baert et al. 2015; Del Bubba et al. 2012; Hager
et al. 2008; Mullen et al. 2003). In Figure 13, we show an ex-
ample of a tetrameric ellagitannin having DOG type linkages
(Karonen et al. 2010)). In the lower part of the structure is a
macrocyclic dimer, oenothein B (1568 Da) which consists of two
tellimagrandin I monomers attached via two m‐DOG type
linkages (2DOG, the blue part of the structure in Figure 13). If
one additional monomer is linked to oenothein B via one DOG
linkage, trimeric oenothein A is formed (MW 2352 Da, the blue
and green parts in Figure 13), and if one more monomer is
linked via one DOG, a tetramer (MW 3136 Da) is formed (the
whole structure in Figure 13). The structure of the tetramer can
FIGURE 11 | Fragmentation pathway of vescalagin in negative ion mode ESI‐MS. The unstable intermediate ions are marked with brackets.
Structures shown are neutral molecules; the m/z values below correspond to the detected ions.
FIGURE 12 | (A) Fragmentation pathway of punicalagin and (B) tentative fragmentation pathway of chebulagic acid in negative ion mode MS.
The m/z values of deprotonated ions corresponding to neutral structures are presented. [Color figure can be viewed at wileyonlinelibrary.com]
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be concluded based on its fragment ions: [M–TIDOG–H]2– atm/z
1175.11, [M–TI2DOG]2– atm/z 1176.12, [M–(TIDOG)2–H]2– atm/z
783.07 and [M–(TIDOGTI2DOG)–H]2– at m/z 784.08 (Karonen
et al. 2010). To minimize isobaric interferences, high‐resolution
MS is needed for the correct identification of these ions (m/z
783, 784 and 785, and m/z 1175 and 1176).
6 | Quantitative Analysis of Tannins
In general, many instrument and sample related aspects need to
be considered in quantitative tannin analysis by mass spec-
trometry. These include sample preparation, optimization of the
ionization and the transmission efficiency of ions, testing of the
effects of sample matrix, and optimization of the calibration
curves taking into account the standardization either by inter-
nal or external standards (Chen et al. 2016). There can be long‐
term signal variations which affect the repeatability and
reproducibility. Quantitative capabilities of mass spectrometry
are improved when chromatographic techniques are used as ion
suppression and contamination effects are reduced and the
effect of sample matrix variability minimized (Chen et al. 2016).
It is also practical to combine MS to UV or diode array detector
as they are nondestructive, reliable and have wide linear range.
In addition, the signal response is not affected by changes in the
mobile phase as long as the eluents are UV transparent at the
wavelengths used. For example, when oligomeric ellagitannins
from willowherb extracts were analyzed with reversed‐phase
UHPLC‐DAD‐ESI‐QqQ‐MS/MS using negative ionization and
the MRM mode, relatively high concentrations of the most
abundant oligomer, that is, dimeric oenothein B, saturated the
detector whereas the scarcer hexamers and heptamers did not
(Baert et al. 2015). Therefore, all extracts were diluted to a range
where oligomers ranging in size from trimers to hexamers were
within the linear range when quantified by MRM and the
dimeric oenothein B was quantified from the UV trace at
280 nm (Baert et al. 2015). This method was selective, sensitive,
and stable with good reproducibility.
For targeted analysis of tannins, triple quadrupoles (QqQ) and
hybrid instruments combining the features of a triple quadru-
pole and a linear ion trap (Q‐Traps) are currently the most used
instruments. In comparison to traditional LC‐MS, LC–MS/MS
provides considerable noise reduction and sensitivity improve-
ments due to enhanced specificity of the SRM/MRM scan
modes. QqQ and Q‐Trap instruments are categorized as low‐
resolution mass spectrometers. Practically, they can measure
them/z ratio of an ion to the nearest integer value (Adrey 2003).
However, different modes of data collection can be utilized,
such as full scan, product ion scan, precursor ion scan, neutral
loss scan, and selected/multiple reaction monitoring. In hybrid
Q‐Trap instruments, the final quadrupole can function as an ion
trap providing MS(n) capabilities and slightly better mass res-
olution when using enhanced MS scan or enhanced product ion
scanning (Sargent 2013).
The main benefit of QqQ and Q‐Trap instruments in tannin
analysis is that several hundred mass transitions can be
FIGURE 13 | The fragmentation of an ellagitannin tetramer in negative ion mode ESI‐MS. On the upper part of the structure, the monomeric
units, that is, tellimagrandin I, are linked to the oligomer via one DOG‐type linkage (TIDOG) and on the lower part of the structure, via two DOG‐type
linkages (TI2DOG). The figure is adapted from Karonen et al. (2010). *The signal is obscured by the signal of the [M–(TIDOGTI2DOG)–H]2– ion. [Color
figure can be viewed at wileyonlinelibrary.com]
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measured during one run when utilizing SRM/MRM method-
ology. In general, in the SRM/MRM analysis of an analyte, the
precursor ion and its most abundant fragment ion are selected
for the quantitative SRM/MRM transition and the second most
abundant fragment ion is used for the qualitative MRM tran-
sition (Baert et al. 2015; Saha and Imran 2020). When the
molecular weight of the tannin exceeds the mass range of the
analyzer, multiply charged ions are utilized (Baert et al. 2015).
To measure hundreds of transitions in one run, the MS/MS
method must be divided into several time‐scheduled windows
with different SRM/MRM channels and time segments
(Guillarme et al. 2010). By focusing a limited number of tran-
sitions within each MRM window, the instrument can allocate
longer dwell times for each transition. For example, Baert et al.
developed an MRM method for the analysis of seven oligomeric
ellagitannins in willow herb leaves: in the method, dwell times
between 16 and 25ms were used in UHPLC‐DAD‐ESI‐QqQ‐
MS/MS to secure at least 12 data points per UHPLC peak for
quantitative work (Baert et al. 2015). On the other hand, when
Lambert et al. analyzed 154 polyphenols in wine, including
flavan‐3‐ols and dimeric PAs, the dwell time of overlapping
analyte peaks were reduced to 5ms (Lambert et al. 2015). By
this, the full capacity of the mass spectrometer was harnessed
and 10−15 data points per analyte peak obtained. The short
dwell time did not yet cause problems but it is worth noticing
that shorter dwell times can lead to lower measurement preci-
sion and sensitivity as the instrument has less time to collect
data for each ion transition. Importantly, to avoid problems
related to retention time variations with UHPLC analyses, one
should always have a proper width in the MRM window
(retention time segmentation), for example, three times the
MRM peak width. If the time windows become very narrow,
small fluctuations outside the original time‐windows might
cause partial loss of detection (Guillarme et al. 2010). Never-
theless, the modern MS/MS instruments can monitor up to 150
simultaneous transitions with enough data points per peak for
quantification purposes (Ehrhardt et al. 2014; Lambert
et al. 2015; Liebler and Zimmerman 2013).
Quantitative studies of tannins using MS/MS are scarce com-
pared to the numerous investigations focused on identification.
At least two challenges may explain this. From a practical point
of view, the absolute tannin quantitation with SRM/MRM is
constrained by the diversity of tannins. Thousands of tannin
structures are found in the plant kingdom while the number of
commercial standards is limited. Compound‐specific SRMs/
MRMs are optimized separately for individual analytes, at least,
by optimizing the cone voltage for the precursor ion and colli-
sion energies for the generation of fragment ions for both
quantitative and qualitative MRM transitions (Tsakalof
et al. 2024). For quantitation, typically the most abundant
fragment ion is selected, while the second most abundant
fragment ion is used to qualitatively ensure accurate identifi-
cation. The ratio between the peak areas corresponding to the
quantitative and qualitative MRM transitions should remain
consistent across different samples and matrices. For method
development, pure compounds should be used whenever pos-
sible. In practice, this requires the isolation and purification of
the reference tannin materials. Although monomers and small
oligomers can be purified and isolated to provide reference
standards (Moilanen et al. 2013; Moilanen and Salminen 2008;
Pfundstein et al. 2010; Saha and Imran 2020; Salminen and
Karonen 2011; Tuominen and Sundman 2013), the purification
of larger oligomers and polymers is difficult due to the number
of isomers and consequent chromatographic behavior of the
larger tannins discussed in Section 4. While method optimiza-
tion can be performed using small amounts of reference stan-
dards, accurate quantification requires purified reference
materials in larger quantities; milligrams are needed for each
tannin. Therefore, studies often utilize relative quantification
and express quantities as equivalents of selected tannins
(Lambert et al. 2015; Pfundstein et al. 2010; Tuominen and
Salminen 2017). Table 1 includes examples of SRM and MRM
transitions used in the literature for tannin quantitation.
For mere screening purposes, the traditional compound‐specific
SRM/MRM methodology faces certain constrains due to two
challenges—individual optimization of MRM transitions and
lack of reference standards, as mentioned above. When
screening thousands of plant samples for their bioactive poly-
phenol content, the primary interest is often in the relative
abundance of polyphenol subgroups rather than the precise
quantification of individual compounds. These kind of data can
be acquired in tannin analysis via qualitative parent ion scan
and neutral loss scan modes (Bai et al. 2022; Robbins
et al. 2014), group‐specific MRMs (Engström et al. 2015, 2014;
Salminen 2018) or more commonly, via chemical
depolymerization or hydrolysis of the tannins before their
UHPLC‐MS/MS analysis (Deshaies et al. 2022; Giribaldi
et al. 2020; Mouls et al. 2011; Singh et al. 2016).
The parent ion and neutral loss scanning methods suffer from
the time needed for data generation and are not suitable for the
simultaneous detection or quantitation of many types of poly-
phenol groups (Engström et al. 2015, 2014; Salminen 2018). In
contrast, the SRM/MRM method has the advantage of short
scanning times as one MRM transition can be measured as fast
as in 5ms, making it especially suitable for the group‐specific
analysis. Traditionally, the SRM/MRM methods of triple
quadrupoles utilize Q1 for precursor selection, Q2 for frag-
mentation, and Q3 for product ion selection, however, many
instruments enable also the use of collision‐induced in‐source
fragmentation by increasing the cone voltage difference
between the sample cone and the extraction cone above a cer-
tain limit (Adrey 2003; de Hoffmann and Stroobant 2001;
Salminen 2018). This in‐source fragmentation technique
allowed the development of group‐specific UHPLC‐MS/MS
methods for tannin analyses (Engström et al. 2015, 2014;
Salminen 2018). Using the methods developed, MRM chro-
matograms serving as fingerprints of HTs and PAs, and other
compounds, such as gallic and quinic acid derivatives, and
quercetin, kaempferol and myricetin glycosides, were recorded
alongside with UV and mass spectra. Thus, an overview of
fingerprints of eight common polyphenol classes was achieved
in a single run. Furthermore, when reference standards are
available, the information can be transformed into quantitative
data (Engström et al. 2015, 2014; Salminen 2018).
The so‐called Engström method relies on the traditional
grouping of tannins based on their chemical structures
(Engström et al. 2014). Compounds in the same polyphenol
group typically share similar or identical building blocks (see
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TABLE 1 | Examples of transitions used in single/multiple reaction monitoring methods in negative ion mode in literature.
Compound
Nominal
mass
Precursor ion
(m/z)a
Product ion
(quant./qual.)
(m/z)a References
Monogalloylglucose 332 331 169/271 Zywicki et al. (2002); Kårlund et al.
(2014); Tuominen and Salminen
(2017); Saha and Imran (2020)
Digalloylglucose 484 483 169/211 Zywicki et al. (2002); Tuominen
and Salminen (2017); Saha and
Imran (2020)
Trigalloylglucose 636 635 169 Zywicki et al. (2002)
635 465/169 Kårlund et al. (2014); Tuominen
and Salminen (2017); Saha and
Imran (2020)
Tetragalloylglucose 788 787 169 Tuominen and Salminen (2017)
787 483/634 Saha and Imran (2020)
Pentagalloylglucose 940 939 169 Tuominen and Salminen (2017)
939 769/617 Lin et al. (2015); Bae et al. (2017);
Saha and Imran (2020)
Hexagalloylglucose 1092 1091 787 Tuominen and Salminen (2017)
1091 939/769 Saha and Imran (2020)
Heptagalloylglucose 1244 621 169 Tuominen and Salminen (2017)
1243 939/1091 Saha and Imran (2020)
Octagalloylglucose 1396 1395 939/769 Saha and Imran (2020)
Tellimagrandin I 786 785 301/275 Kårlund et al. (2014); Saha and
Imran (2020)
Tellimagrandin II 938 937 301 Tuominen and Salminen (2017)
Pedunculagin 784 783 301/275 Kårlund et al. (2014); Saha and
Imran (2020)
2,3‐(S)‐HHDP‐glucose 482 481 301 Kårlund et al. (2014)
Strictinin 634 633 301 Saha and Imran (2020)
Galloyl‐HHDP‐glucose 634 633 301 Kårlund et al. (2014)
Casuarictin 936 935 633/301 Gasperotti et al. (2015)
Vescalagin 934 933 301 García‐Estévez et al. (2012);
Engström et al. (2015)
933 915/301 Saha and Imran (2020)
466 301 Stark et al. (2010)
Castalagin 934 933 301/425 Saha and Imran (2020)
933 631 García‐Estévez et al. (2012)
466 301 Stark et al. (2010)
Grandinin/roburin E 1066 1065 249 García‐Estévez et al. (2012)
Vescavaloninic acid 1102 1101 1083/569 Saha and Imran (2020)
Castavaloninic acid 1102 1101 1057/425 Saha and Imran (2020)
Acutissimin A/B 1206 602 457 Stark et al. (2010)
Epiacutissimin A/B 1206 602 457 Stark et al. (2010)
Geraniin 952 951 933 Tuominen and Salminen (2017)
951 301/933 Saha and Imran (2020)
Carpinusin 952 951 933 Tuominen and Salminen (2017)
(Continues)
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TABLE 1 | (Continued)
Compound
Nominal
mass
Precursor ion
(m/z)a
Product ion
(quant./qual.)
(m/z)a References
Geraniic acid 952 907 291 Tuominen and Salminen (2017)
Ascorgeraniin 1110 1109 933 Tuominen and Salminen (2017)
1109 301/973 Saha and Imran (2020)
Sylvatiin A 1144 571 169 Tuominen and Salminen (2017)
Sylvatiin B 992 495 169 Tuominen and Salminen (2017)
Sylvatiin C 1348 673 169 Tuominen and Salminen (2017)
Sylvatiin D 1160 579 169 Tuominen and Salminen (2017)
Sylvatiin E 840 839 373 Tuominen and Salminen (2017)
Chebulagic acid 954 953 301/275 Kumar et al. (2017); Tuominen and
Salminen (2017); Saha and
Imran (2020)
Chebulanin 652 651 169/481 Saha and Imran (2020)
Chebulinic acid 956 955 169/205 Saha and Imran (2020)
955 337/617 Kumar et al. (2017)
Oenothein B 1568 783 765/301 Baert et al. (2015); Engström
et al. (2015)
Oenothein A 2352 1176 301/275 Baert et al. (2015)
Tetrameric ET 3136 1045 301/275 Baert et al. (2015)
Pentameric ET 3920 1306 301/275 Baert et al. (2015)
Hexameric ET 4704 1568 301/275 Baert et al. (2015)
Heptameric ET 5488 1829 275/301 Baert et al. (2015)
Sanguiin H6 1870 934 633/301 Gasperotti et al. (2015)
934 301/275
Gemin A 1872 935 301/275 Saha and Imran (2020)
Agrimoniin 1870 934 633/301 Gasperotti et al. (2015)
934 301 Kårlund et al. (2014)
Lambertianin C 2748 935 301/275 Saha and Imran (2020)
Cocciferin D2 1868 933 301/275 Saha and Imran (2020)
Rubusuaviin C 2804 934 301/275 Saha and Imran (2020
Salicarinin A 1868 933 249/301 Saha and Imran (2020)
Salicarinin B 1868 933 301/249 Saha and Imran (2020)
Rugosin D 1874 935 301/275 Saha and Imran (2020)
Rugosin E 1722 860 301/275 Saha and Imran (2020)
Proanthocyanidin B1 578 577 289/407/125 Delgado de la Torre et al. (2013);
Ehrhardt et al. (2014)
577 425/407 Margalef et al. (2014)
Proanthocyanidin B2 578 577 289/407/125 Delgado de la Torre et al. (2013);
Ehrhardt et al. (2014)
577 289/425 Ortega et al. (2010)
577 425/407 Margalef et al. (2014)
Proanthocyanidin B3 578 577 289/407 Ehrhardt et al. (2014)
577 425/407 Margalef et al. (2014)
Proanthocyanidin A2 576 575 285/539/449 Delgado de la Torre et al. (2013)
(Continues)
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Figure 1). Regarding tannins, all galloylglucoses and gallo-
tannins yield a gallic acid fragment ion at m/z 169, see
Figure 10) that could be detected by the parent ion scan of m/z
169 in negative ion mode (Engström et al. 2015; Salminen 2018).
Similarly, all hexahydroxydiphenoyl‐containing ellagitannins
yielded a fragment ion at m/z 301 (see Figure 11). When these
ions were selected in Q1 as precursor ions, followed by frag-
mentation in Q2 and subsequent qualifier and quantifier
selection in Q3, group‐specific MRM methods were produced
for HTs (Engström et al. 2015).
The need for rapid screening methods for PAs is even more
urgent due to the chromatographic challenges and poor ion-
ization of the larger PAs in their analysis via LC‐MS (see
Section 4). To facilitate the rapid analysis of PAs, a method was
developed to separately measure the procyanidin and prodel-
phinidin units in the PA structure (Engström et al. 2014).
Conceptually, the method is similar to the widely adopted
thiolysis and phloroglucinolysis methods (Thompson
et al. 1972; Hemingway 1989b; Matthews et al. 1997; Fulcrand
et al. 1999; Guyot et al. 2001; Kennedy and Jones 2001), which
are used to depolymerize PAs into smaller units for analysis.
However, instead of using chemical reactions for the
depolymerization of PAs, this method fragments them in the
ESI interface of a mass spectrometer. The quinone‐methide
fragmentation in the ion‐source allows determining the pro-
cyanidin/prodelphinidin ratios and the mean degree of polym-
erization for the PAs present in any studied sample. The most
powerful feature of the method is that the degradation of the
PAs is done after the chromatographic step. This is beneficial as
the original compound profile is retained in the form of chro-
matographic fingerprints (Engström et al. 2015, 2014;
Salminen 2018). Consequently, it allows not only qualitative but
also quantitative comparison via sample‐specific fingerprints of
the PA composition, procyanidin/prodelphinidin ratio and
mean degree of polymerization throughout the chromato-
graphic hump produced by the larger PA oligomers and poly-
mers (Engström et al. 2014; Salminen 2018). The group‐specific
methodology can be applied also for other compound groups if
they share a common structural unity. For example, Laitila
et al. developed a group‐specific method for the analysis of
anthocyanins and anthocyanin adducts in red wines, which
utilizes the same in‐source fragmentation approach (Laitila
et al. 2019).
7 | Bioactivities of Tannins by Mass
Spectrometric Tools
In addition to the qualitative and quantitative analysis of tan-
nins as such, MS can be used as a tool for bioactivity studies.
When a studied plant species expresses certain bioactivity, the
next step is to understand the underlying mechanisms and what
type of compounds are the most active and why. In plants, the
protective role of tannins against herbivory and many
pathogens actualizes through two main mechanisms, protein
binding and oxidation reactions (Appel 1993; Salminen and
Karonen 2011). In addition to the plant–herbivore interplay,
tannin–protein interactions are important in numerous other
plant related domains and the interest in studying the com-
plexes between tannins and proteins has increased steadily
during the past few decades. Mass spectrometry has been
widely used to study tannin–protein complexes and to under-
stand the exact roles of tannins and proteins in these
TABLE 1 | (Continued)
Compound
Nominal
mass
Precursor ion
(m/z)a
Product ion
(quant./qual.)
(m/z)a References
A‐type procyanidin dimer 576 575 285 Pérez‐Jiménez and Torres (2012)
B‐type prodelphinidin dimer 610 609 125 Tuominen and Salminen (2017)
A‐type prodelphinidin dimer
(PD‐PC)
592 591 465 Pérez‐Jiménez and Torres (2012)
B‐type propelargonidin
dimer (PP‐PC)
562 561 289 Pérez‐Jiménez and Torres (2012)
B‐type procyanidin trimer 866 865 289 Kårlund et al. (2014)
865 577/695 Ortega et al. (2010); Pérez‐Jiménez
and Torres (2012)
B‐type propelargonidin
trimer
850 849 577 Pérez‐Jiménez and Torres (2012)
PA tetramer 1154 1153 865/575 Ortega et al. (2010)
PA pentamer 1442 1441 1028/1151 Ortega et al. (2010)
PA hexamer 1730 1729 1153/863 Ortega et al. (2010)
PA heptamer 2018 1008 865/575 Ortega et al. (2010)
PA octamer 2306 1152 875/983 Ortega et al. (2010)
PA nonamer 2594 1296 577/1152 Ortega et al. (2010)
Abbreviations: ET = ellagitannin, HHDP= hexahydroxydiphenoyl, PA= proantocyanidin, PC = procyanidin, PD = prodelphinidin, PP = propelargonidin
aValues with decimals were rounded to unit mass.
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interactions (Baron et al. 2019; Canon et al. 2015, 2011, 2009; Di
et al. 2024; Dias et al. 2016; Engström, Sun et al. 2016; Ghahri
et al. 2021; Manjón et al. 2024; Mau et al. 2011; Sarni‐Manchado
and Cheynier 2002; Shraberg et al. 2015; Simon et al. 2003). To
explain the mechanisms behind tannin‐protein interactions and
to understand what types of tannins are the most active, pure
compounds must be utilized in the activity tests.
While the protein precipitation capacity of tannins, that is, ability
to precipitate proteins and other macromolecules out of aqueous
solution, has traditionally been recognized as the source of
defensive function against herbivory, oxidation and the following
reactions are more prevalent at high pH (Appel 1993; Gross
et al. 2008; Johnson and V. Barbehenn 2000; Kim et al. 2018;
Salminen and Karonen 2011). Tannins can be oxidized even at
low to neutral pH by plant polyphenol oxidases and peroxidases
(Appel 1993; Kim et al. 2018; Salminen and Karonen 2011). LC‐
MS has been used to study the oxidation of both purified tannins
and tannins present in plant extracts to explore the structure–
activity relationships (Kim et al. 2020, 2018; Tuominen and
Sundman 2013; Vihakas et al. 2014). Moreover, the in vitro oxi-
dation of plant tannins in insect herbivores has been examined
by LC‐MS (Salminen and Lempa 2002; Vihakas et al. 2015). In
these applications, the sensitivity of LC‐MS and LC‐MS/MS is
needed due to the small amounts of sample materials and low
tannin concentrations, where less sensitive methods, such as LC‐
DAD, are inadequate for quantifying all compounds present in
the sample.
The structural changes of HTs caused by oxidation on a
molecular level can be revealed by NMR spectroscopy (Hatano
et al. 2004; Tanaka et al. 1990) but its use is limited to stable
oxidation products that can be isolated in large quantities (mg
scale). LC‐MS and especially LC‐DAD‐MS, are effective tools to
study the oxidative activity and kinetics of oxidation of purified
HTs (Tuominen and Sundman 2013). These approaches also
allow the tentative identification of the main degradation
products based on UV and MS spectra without isolating the
compounds. Regarding PAs, their oxidation reactions are gen-
erally well‐known but site of modification has been reported
mainly for individual smaller PAs (Burger et al. 1990; Ferreira
et al. 1992; Guyot et al. 1996; Hibi and Yanase 2019; Laks
et al. 1987). Earlier, the definitive structural studies of PAs and
their oxidation products often relied on the use of NMR (Burger
et al. 1990; Ferreira et al. 1992; Laks et al. 1987). Lately, MS has
been successfully applied for the analysis of PA oxidation (Chen
et al. 2014; Imran et al. 2021; Karonen et al. 2021; Kondo
et al. 2000; Mouls and Fulcrand 2012). These studies showed
different reactivities for procyanidins and prodelphinidins,
suggesting that the presence of prodelphinidin units enhanced
the probability of modification reactions. By high‐resolution MS
and by measuring qualitative changes in the procyanidin/pro-
delphinidin ratio and mean degree of polymerization by MRM
methods, the main reaction routes for both A‐ and B‐type PAs
were concluded to be intramolecular (Imran et al. 2021;
Karonen et al. 2021). For prodelphinidin‐rich and galloylated
PAs, also intermolecular reactions were indicated (Imran
et al. 2021; Karonen et al. 2021).
One special field where MS is often used to study tannin oxi-
dation is the studies related to wine production. During wine
aging, the controlled oxygen levels allow the wine to evolve,
particularly via the chemical modification of the tannins, for
example, via oxidation of tannins and the following inter‐ and
intramolecular reactions (Arapitsas et al. 2014; Petit et al. 2019).
Tannins and their oxidation products are important for the
sensory quality of the wine; they are thus responsible for
astringency, bitterness and contribute to its color. The ability to
measure the oxidative evolution of the wine tannins enables to
predict the optimal aging times when linked to the modulation
of astringency feeling (Deshaies et al. 2022). Often untargeted
approaches are used to evaluate the overall changes of tannins
in the wines (Arapitsas et al. 2020; Arbulu et al. 2015; Castro
et al. 2014) but alternatively targeted analysis utilizing marker
ions for the quantification of specific changes can be used
(Deshaies et al. 2022; Mouls and Fulcrand 2012; Ontañón
et al. 2020). In addition, the changes can be measured indirectly
by determining the disappearance of the original PAs. All these
approaches face the same constrains as PA analysis in general:
the chromatographic co‐elution of the numerous isomers cre-
ates the characteristic chromatographic hump and the effi-
ciency of ionization is reduced by the saturation of more
abundant PAs (Mouls et al. 2011). Moreover, in comparison to
intact tannins, their oxidation further increases the chemical
diversity and structural complexity making their identification
challenging (Imran et al. 2021; Karonen et al. 2021; Mouls and
Fulcrand 2012).
Chemical depolymerization of the PAs to their monomeric units
are often applied to measure the changes in their concentration
and mean degree of polymerization. This is especially beneficial
when searching for specific oxidation markers. The new
covalent bonds formed in oxidized PA structures are not de-
polymerized efficiently, as oxidation makes the initial PAs
resistant to the depolymerization conditions (Mouls and
Fulcrand 2015, 2012). Thus, dimeric and trimeric oxidation
markers are observed after chemical depolymerization of the
tannins (Deshaies et al. 2022) and these oxidation markers can
be used to evaluate the oxidation state of the wine, for example,
for quality control purposes. However, these chemical
depolymerization methods do not enable characterization of the
changes in the overall profiles of the oligomeric and polymeric
PAs as the information on individual PA oligomers or polymers
and their oxidized products are lost. To avoid this, Imran et al.
utilized a group‐specific approach to study the oxidation and
stability of different types of natural PAs via nonspecific alka-
line oxidation (Imran et al. 2021). This approach allowed
quantifying the decrease in PA concentration and changes
in PA composition as mentioned above. However, there is still a
need for new analytical method developments in the identifi-
cation and characterization of the oxidation products of PAs by
MS. Many of the oxidation products are not detected under the
standard ESI‐MS conditions even though a hump indicating
PAs is detected by UV at 280 nm (Deshaies et al. 2022; Karonen
et al. 2021).
The ability of tannins to interact with proteins in biological
systems is important for many plant related domains, including
the most basic functions in plant physiology and ecology, but
also increasingly in applications related to agriculture, foods/
feeds and medicine. Due to the increasing interest in these in-
teractions, their consequences, and their utilization in various
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applications, numerous physicochemical methods have been
developed over the years to study, for example, complex for-
mation, protein precipitation, binding affinities, binding stoi-
chiometry, conformational changes, and kinetics (Baron
et al. 2019; Canon et al. 2015, 2011, 2009; Di et al. 2024; Dias
et al. 2016; Engström et al. 2022, 2019; Engström, Sun
et al. 2016; Ghahri et al. 2021; Mau et al. 2011; Sarni‐Manchado
and Cheynier 2002; Shraberg et al. 2015; Simon et al. 2003).
MS has many benefits when studying tannin−protein complexes.
The underlying reasons are the same as those mentioned above
for other applications: it is a sensitive technique, and therefore,
only small sample volumes are required. In addition, it allows the
determination of the stoichiometry of the formed complexes, the
tentative mechanism of complex formation and the structures of
the complexes formed (Baron et al. 2019; Canon
et al. 2015, 2011, 2009; Di et al. 2024; Dias et al. 2016; Engström,
Sun et al. 2016; Ghahri et al. 2021; Mau et al. 2011; Nguyen
et al. 2021; Sarni‐Manchado and Cheynier 2002; Shraberg
et al. 2015; Simon et al. 2003). The main challenge in MS analysis
is the stability of the noncovalent tannin−protein complexes.
Partially due to this, soft ionization techniques MALDI
(Engström, Sun et al. 2016; Ghahri et al. 2021; Mau et al. 2011;
Simon et al. 2003) and ESI (Baron et al. 2019; Canon
et al. 2015, 2011; Dias et al. 2016; Sarni‐Manchado and
Cheynier 2002; Shraberg et al. 2015) are the most used ioniza-
tion techniques. The simplicity of MALDI benefits data inter-
pretation as only singly charged ions are present in the mass
spectra of the tannin–protein complexes as shown in Figure 14
(Mané et al. 2007). However, MALDI may suffer from low
sample‐to‐sample and scan‐to‐scan reproducibility and the sig-
nal level strongly depends on the laser beam homogeneity and
irradiance, sample preparation (i.e., the choice of a suitable
matrix and the solvents), the analyte properties, sample purity,
as well as the crystallization process of the sample (Gusev
et al. 1995). In addition, neither ESI nor MALDI can
unambiguously describe the tannin−protein complexes as
noncovalent or covalent, which is often a primary interest
(Engström, Sun et al. 2016). In ESI, tentative identification of
complex types and evaluation of complex strengths can be
achieved by increasing the ionization voltage, which promotes
dissociation of noncovalent complexes in the gas phase through
ion–molecule collisions at the atmospheric–vacuum interface.
However, this is also a source of misinterpretation, as detecting
complexes requires optimizing ionization to increase sensitivity
while minimizing the dissociation of noncovalent complexes
(Canon et al. 2009; Sarni‐Manchado and Cheynier 2002). Also,
as the complexes are analyzed in the gas phase, the results
cannot be directly reflected to the reactions in solution
(Loo 1997; Sarni‐Manchado and Cheynier 2002).
The analyzers used for detection of tannin–protein complexes
are variable including quadrupoles (Q and QqQ), TOFs, quad-
rupole ion traps, orbitraps, FT‐ICR instruments, and different
hybrid mass analyzers (Baron et al. 2019; Canon et al. 2015;
Dias et al. 2016; Engström, Sun et al. 2016; Perez‐Gregorio
et al. 2014; Sarni‐Manchado and Cheynier 2002; Shraberg
et al. 2015; Vergé et al. 2002). Also, ion mobility spectrometry
has been successfully used to study the interactions between
human salivary proline‐rich proteins and the galloylated flavan‐
3‐ol epigallocatechin gallate (Canon et al. 2011). It offers the
significant advantage of enabling the determination of complex
distributions and the structural isomers of each species (Bowers
et al. 1993; Niu et al. 2013). While other mass spectrometric
methods evaluate tannin–protein complexation and the result-
ing complexes as a static process, ion mobility spectrometry
enables the assessment of complexation as a dynamic process,
in which conformational changes in tannin and protein struc-
tures play a key role in the observed function or activity (Canon
et al. 2011).
8 | Conclusions
Various mass spectrometric techniques provide powerful tools
for the characterization and quantification of tannins. In the
qualitative analysis of diverse oligo‐ and polymeric tannins, ESI
combined with high‐resolution mass analyzers is particularly
effective, as ESI enables the ionization of high‐molecular‐
weight tannins in the form of multiply charged ions, thereby
significantly expanding the accessible mass range. For quanti-
tative tannin analysis, tandem MS techniques and MRM stand
out due to their superior specificity and sensitivity. These
methods can also provide structural information, such as the
procyanidin/prodelphinidin ratio and mean degree of polym-
erization for PAs, as well as the presence of galloyl and hex-
ahydroxy groups in HT structures. Mass spectrometry is also
well‐suited for investigating the bioactivities of tannins, partic-
ularly in elucidating tannin–protein interactions. Future ad-
vancements in MS technologies, including the integration of
conventional ionization methods with novel ionization pro-
cesses, may offer enhanced capabilities for tannin imaging and
direct analysis in biological matrices. These developments could
also lead to improved tools for screening the bioactivities and
other beneficial effects of tannins. Moreover, automated ana-
lytical platforms may streamline practical tannin analysis, while
advanced data processing approaches, such as molecular
FIGURE 14 | Example of data produced with MALDI‐TOF in pos-
itive ion mode when bovine serum albumin (average MW 66.5 kDa) and
oligomeric ellagitannin oenothein A (MW 2353.6 Da) were incubated at
pH 5 (black color) or pH 6.7 (orange color), and after removal of
unstable complexes by ultrafiltration (pH 5 turquoise, pH 6.7 blue).
Figure modified from (Engström, Sun et al. 2016). [Color figure can be
viewed at wileyonlinelibrary.com]
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networking and machine learning, can assist in the laborious
and time‐consuming interpretation of complex MS data.
Author Contributions
Marica T. Engström: conceptualization, data curation, investigation,
resources, software, visualization, writing – original draft, writing –
review and editing.Maarit Karonen: conceptualization, data curation,
investigation, resources, software, visualization, writing – original draft,
writing – review and editing.
Acknowledgments
The authors thank professor Juha‐Pekka Salminen for reviewing and
commenting the first draft of the paper. All current and previous NCRG
members are acknowledged for sharing the passion on tannin chemistry
and the fruitful discussions during the years. Open access publishing
facilitated by Turun yliopisto, as part of the Wiley ‐ FinELib agreement.
References
Adamson, G. E., S. A. Lazarus, A. E. Mitchell, et al. 1999. “Hplc Method
for the Quantification of Procyanidins in Cocoa and Chocolate Samples
and Correlation to Total Antioxidant Capacity.” Journal of Agricultural
and Food Chemistry 47: 4184–4188. https://doi.org/10.1021/jf990317m.
Adrey, R. E. 2003. Liquid Chromatography‐Mass Spectrometry: An
Introduction. John Wiley & Sons Ltd.
Aguilar‐Zárate, P., J. E. Wong‐Paz, M. Michel, et al. 2017. “Character-
isation of Pomegranate‐Husk Polyphenols and Semi‐Preparative Frac-
tionation of Punicalagin.” Phytochemical Analysis: PCA 28: 433–438.
https://doi.org/10.1002/pca.2691.
Appel, H. M. 1993. “Phenolics in Ecological Interactions: The Impor-
tance of Oxidation.” Journal of Chemical Ecology 19: 1521–1552. https://
doi.org/10.1007/BF00984895.
Arapitsas, P., G. Speri, A. Angeli, D. Perenzoni, and F. Mattivi. 2014.
“The Influence of Storage on the “Chemical Age” of Red Wines.”
Metabolomics 10: 816–832. https://doi.org/10.1007/s11306-014-0638-x.
Arapitsas, P., M. Ugliano, M. Marangon, et al. 2020. “Use of Untargeted
Liquid Chromatography‐Mass Spectrometry Metabolome to Discrimi-
nate Italian Monovarietal Red Wines, Produced in Their Different
Terroirs.” Journal of Agricultural and Food Chemistry 68: 13353–13366.
https://doi.org/10.1021/acs.jafc.0c00879.
Arbulu, M., M. C. Sampedro, A. Gómez‐Caballero, M. A. Goicolea, and
R. J. Barrio. 2015. “Untargeted Metabolomic Analysis Using Liquid
Chromatography Quadrupole Time‐of‐Flight Mass Spectrometry for
Non‐Volatile Profiling of Wines.” Analytica Chimica Acta 858: 32–41.
https://doi.org/10.1016/j.aca.2014.12.028.
Bae, S., S. Y. Kim, M. H. Do, C. H. Lee, and Y.‐J. Song. 2017. “1,2,3,4,6‐
Penta‐O‐Galloyl‐ß‐D‐Glucose, a Bioactive Compound in Elaeocarpus
sylvestris Extract, Inhibits Varicella‐Zoster Virus Replication.” Antiviral
Research 144: 266–272. https://doi.org/10.1016/j.antiviral.2017.06.018.
Baert, N., M. Karonen, and J.‐P. Salminen. 2015. “Isolation, Char-
acterisation and Quantification of the Main Oligomeric Macrocyclic
Ellagitannins in Epilobium Angustifolium by Ultra‐High Performance
Chromatography With Diode Array Detection and Electrospray Tan-
dem Mass Spectrometry.” Journal of Chromatography A 1419: 26–36.
https://doi.org/10.1016/j.chroma.2015.09.050.
Bai, Z., R. Yu, T. Zheng, et al. 2022. “A Novel Strategy for Unveiling
Spatial Distribution Pattern of Gallotannins in Paeonia rockii and
Paeonia ostii Based on LC–QTRAP–MS.” Metabolites 12: 326. https://
doi.org/10.3390/metabo12040326.
Barofsky, D. F. 1989. “FAB‐MS Applications in the Structure Elucida-
tion of Proanthocyanidin Structures.” In Chemistry and Significance of
Condensed Tannins, edited by R. W. Hemingway and J. J. Karchesy,
175–195. J. J. Plenum Press.
Baron, G., A. Altomare, L. Fumagalli, et al. 2019. “Development of a
Direct ESI‐MS Method for Measuring the Tannin Precipitation Effect of
Proline‐Rich Peptides and In Silico Studies on the Proline Role in
Tannin‐Protein Interactions.” Fitoterapia 136: 104163. https://doi.org/
10.1016/j.fitote.2019.05.002.
Barry, K. M., N. W. Davies, and C. L. Mohammed. 2001. “Identification
of Hydrolysable Tannins in the Reaction Zone of Eucalyptus nitens
Wood by High Performance Liquid Chromatography‐Electrospray Io-
nisation Mass Spectrometry.” Phytochemical Analysis 12: 120–127.
https://doi.org/10.1002/pca.548.
Behrens, A., N. Maie, H. Knicker, and I. Kögel‐Knabner. 2003.
“MALDI‐TOF Mass Spectrometry and PSD Fragmentation as Means for
the Analysis of Condensed Tannins in Plant Leaves and Needles.”
Phytochemistry 62: 1159–1170. https://doi.org/10.1016/S0031-9422(02)
00660-X.
Berardini, N., R. Carle, and A. Schieber. 2004. “Characterization of
Gallotannins and Benzophenone Derivatives From Mango (Mangifera
indica L. cv. ‘Tommy Atkins’) Peels, Pulp and Kernels by High‐
Performance Liquid Chromatography/Electrospray Ionization Mass
Spectrometry.” Rapid Communications in Mass Spectrometry: RCM 18:
2208–2216. https://doi.org/10.1002/rcm.1611.
Bowers, J. J., H. P. Gunawardena, A. Cornu, et al. 2018. “Rapid
Screening of Ellagitannins in Natural Sources via Targeted Reporter Ion
Triggered Tandem Mass Spectrometry.” Scientific Reports 8: 10399.
https://doi.org/10.1038/s41598-018-27708-3.
Bowers, M. T., P. R. Kemper, G. Von Helden, and P. A. M. Van Koppen.
1993. “Gas‐Phase Ion Chromatography: Transition Metal State Selection
and Carbon Cluster Formation.” Science 260, no. 1979: 1446–1451.
https://doi.org/10.1126/science.260.5113.1446.
Burger, J. F. W., H. Kolodziej, R. W. Hemingway, J. P. Steynberg,
D. A. Young, and D. Ferreira. 1990. “Oligomeric Flavanoids. Part W.
Base‐Catalyzed Pyran Rearrangements of Procyanidin B‐2, and Evi-
dence for the Oxidative Transformation of B‐ to A‐Type Procyanidins.”
Tetrahedron 46: 5733–5740.
Canon, F., R. Ballivian, F. Chirot, et al. 2011. “Folding of a Salivary
Intrinsically Disordered Protein Upon Binding to Tannins.” Journal of
the American Chemical Society 133: 7847–7852. https://doi.org/10.1021/
ja200534f.
Canon, F., F. Paté, E. Meudec, et al. 2009. “Characterization, Stoichi-
ometry, and Stability of Salivary Protein‐Tannin Complexes by ESI‐MS
and ESI‐MS/MS.” Analytical and Bioanalytical Chemistry 395:
2535–2545. https://doi.org/10.1007/s00216-009-3180-3.
Canon, F., S. Ployon, J.‐P. Mazauric, et al. 2015. “Binding Site of Dif-
ferent Tannins on a Human Salivary Proline‐Rich Protein Evidenced by
Dissociative Photoionization Tandem Mass Spectrometry.” Tetrahedron
71: 3039–3044. https://doi.org/10.1016/j.tet.2014.11.013.
Castro, C. C., R. C. Martins, J. A. Teixeira, and A. C. Silva Ferreira. 2014.
“Application of a High‐Throughput Process Analytical Technology
Metabolomics Pipeline to Port Wine Forced Ageing Process.” Food
Chemistry 143: 384–391. https://doi.org/10.1016/j.foodchem.2013.
07.138.
Chen, L., P. Yuan, K. Chen, Q. Jia, and Y. Li. 2014. “Oxidative Con-
version of B‐ to A‐Type Procyanidin Trimer: Evidence for Quinone
Methide Mechanism.” Food Chemistry 154: 315–322.
Chen, Y. C., P. L. Urban, and Y. S. Wang. 2016. Time‐Resolved Mass
Spectrometry: From Concept to Applications. John Wiley & Sons
Incorporated.
Cheynier, V., and H. Fulcrand. 2003. “Analysis of Polymeric Proan-
thocyanidins and Complex Polyphenols.” In Methods in Polyphenol
Analysis, edited by C. Santos‐Buelga and G. Williamson. The Royal
Society of Chemistry.
20 Mass Spectrometry Reviews, 2025
 10982787, 0, D
ow
nloaded from
 https://analyticalsciencejournals.onlinelibrary.wiley.com/doi/10.1002/mas.70013 by University of Turku, W
iley Online Library on [28/11/2025]. See the Terms and Conditions (https://onlinelibrary.wiley.com/terms-and-conditions) on W
iley Online Library for rules of use; OA articles are governed by the applicable Creative Commons License
Cobo, A., A. Alejo‐Armijo, D. Cruz, J. Altarejos, S. Salido, and E. Ortega
‐Morente. 2024. “Halogenated Analogs to Natural A‐Type Proantho-
cyanidins: Evaluation of Their Antioxidant and Antimicrobial Proper-
ties and Possible Application in Food Industries.” Molecules 29: 3622.
https://doi.org/10.3390/molecules29153622.
Cooper, H. J., and A. G. Marshall. 2001. “Electrospray Ionization Fou-
rier Transform Mass Spectrometric Analysis of Wine.” Journal of
Agricultural and Food Chemistry 49: 5710–5718.
de Hoffmann, E., and V. Stroobant. 2001. Mass Spectrometry: Principles
and Applications. John Wiley & Sons Ltd.
de Pascual‐Teresa, S., Y. Gutiérrez‐Fernández, J. C. Rivas‐Gonzalo,
and C. Santos‐Buelga. 1998. “Characterization of Monomeric and
Oligomeric Flavan‐3‐ols From Unripe Almond Fruits.” Phytochemical
Analysis 9: 21–27.
de Pascual‐Teresa, S., J. C. Rivas‐Gonzalo, and C. Santos‐Buelga. 2000.
“Prodelphinidins and Related Flavanols in Wine.” International Journal
of Food Science and Technology 35: 33–40.
Deffieux, D., A. Natangelo, G. Malik, L. Pouységu, J. Charris, and
S. Quideau. 2011. “First and Biomimetic Total Synthesis of a Member of
the C‐Glucosidic Subclass of Ellagitannins, 5‐O‐Desgalloylepipunicacortein
A.” Chemical Communications 47: 1628–1630. https://doi.org/10.1039/
c0cc04007j.
Del Bubba, M., L. Checchini, U. Chiuminatto, S. Doumett, D. Fibbi, and
E. Giordani. 2012. “Liquid Chromatographic/Electrospray Ionization
Tandem Mass Spectrometric Study of Polyphenolic Composition of
Four Cultivars of Fragaria vesca L. Berries and Their Comparative
Evaluation.” Journal of Mass Spectrometry 47: 1207–1220. https://doi.
org/10.1002/jms.3030.
Delgado de La Torre, M. P., C. Ferreiro‐Vera, F. Priego‐Capote, and
M. D. Luque de Castro. 2013. “Anthocyanidins, Proanthocyanidins, and
Anthocyanins Profiling in Wine Lees by Solid‐Phase Extraction‐Liquid
Chromatography Coupled to Electrospray Ionization Tandem Mass
Spectrometry With Data‐Dependent Methods.” Journal of Agricultural
and Food Chemistry 61: 12539–12548. https://doi.org/10.1021/
jf404194q.
Deshaies, S., F. Garcia, L. Suc, C. Saucier, and L. Mouls. 2022. “Study of
the Oxidative Evolution of Tannins During Syrah Red Wines Ageing by
Tandem Mass Spectrometry.” Food Chemistry 385: 132538. https://doi.
org/10.1016/j.foodchem.2022.132538.
Di, X., Y. Li, X. Qin, Q. Wang, and G. Liu. 2024. “Investigating the Effect
of Whey Protein Isolate: Proanthocyanidin Complex Ratio on the Sta-
bility and Antioxidant Capacity of Pickering Emulsions.” International
Journal of Biological Macromolecules 279: 135342. https://doi.org/10.
1016/j.ijbiomac.2024.135342.
Dias, R., M. R. Perez‐Gregorio, N. Mateus, and V. De Freitas. 2016.
“Interaction Study Between Wheat‐Derived Peptides and Procyanidin
B3 by Mass Spectrometry.” Food Chemistry 194: 1304–1312. https://doi.
org/10.1016/j.foodchem.2015.08.108.
Duckstein, S. M., E. M. Lotter, U. Meyer, U. Lindequist, and
F. C. Stintzing. 2013. “Phenolic Constituents From Alchemilla Vulgaris
L. and Alchemilla mollis (Buser) Rothm. at Different Dates of Harvest.”
Zeitschrift fur Naturforschung. C, Journal of Biosciences 68: 529–540.
https://doi.org/10.5560/znc.2012.67c0529.
Ehrhardt, C., P. Arapitsas, M. Stefanini, G. Flick, and F. Mattivi. 2014.
“Analysis of the Phenolic Composition of Fungus‐Resistant Grape
Varieties Cultivated in Italy and Germany Using UHPLC‐MS/MS.”
Journal of Mass Spectrometry 49: 860–869. https://doi.org/10.1002/
jms.3440.
Engström, M. T., J. Arvola, S. Nenonen, et al. 2019. “Structural
Features of Hydrolyzable Tannins Determine Their Ability to Form
Insoluble Complexes With Bovine Serum Albumin.” Journal of
Agricultural and Food Chemistry 67: 6798–6808. https://doi.org/10.
1021/acs.jafc.9b02188.
Engström, M. T., M. Karonen, J. R. Ahern, et al. 2016. “Chemical
Structures of Plant Hydrolyzable Tannins Reveal Their In Vitro Activity
Against Egg Hatching and Motility of Haemonchus contortus Nema-
todes.” Journal of Agricultural and Food Chemistry 64: 840–851. https://
doi.org/10.1021/acs.jafc.5b05691.
Engström, M. T., M. Pälijärvi, C. Fryganas, J. H. Grabber, I. Mueller‐
Harvey, and J.‐P. Salminen. 2014. “Rapid Qualitative and Quantitative
Analyses of Proanthocyanidin Oligomers and Polymers by UPLC‐MS/
MS.” Journal of Agricultural and Food Chemistry 62: 3390–3399. https://
doi.org/10.1021/jf500745y.
Engström, M. T., M. Pälijärvi, and J.‐P. Salminen. 2015. “Rapid Fin-
gerprint Analysis of Plant Extracts for Ellagitannins, Gallic Acid, and
Quinic Acid Derivatives and Quercetin‐, Kaempferol‐ and Myricetin‐
Based Flavonol Glycosides by UPLC‐QQQ‐MS/MS.” Journal of
Agricultural and Food Chemistry 63: 4068–4079. https://doi.org/10.
1021/acs.jafc.5b00595.
Engström, M. T., X. Sun, M. P. Suber, M. Li, J.‐P. Salminen, and
A. E. Hagerman. 2016. “The Oxidative Activity of Ellagitannins Dictates
Their Tendency to Form Highly Stabilized Complexes With Bovine
Serum Albumin at Increased pH.” Journal of Agricultural and Food
Chemistry 64: 8994–9003. https://doi.org/10.1021/acs.jafc.6b01571.
Engström, M. T., V. Virtanen, and J.‐P. Salminen. 2022. “Influence of
the Hydrolyzable Tannin Structure on the Characteristics of Insoluble
Hydrolyzable Tannin‐Protein Complexes.” Journal of Agricultural and
Food Chemistry 70: 13036–13048. https://doi.org/10.1021/acs.jafc.
2c01765.
Escribano‐Bailon, T., Y. Gutierrez‐Fernandez, J. C. Rivas‐Gonzalo, and
C. Santos‐Buelga. 1992. “Characterization of Procyanidins of Vitis
vinifera Variety Tinta del Pais Grape Seeds.” Journal of Agricultural and
Food Chemistry 40: 1794–1799. https://doi.org/10.1021/jf00022a013.
Es‐Safi, N.‐E., S. Guyot, and P.‐H. Ducrot. 2006. “NMR, ESI/MS, and
MALDI‐TOF/MS Analysis of Pear Juice Polymeric Proanthocyanidins
With Potent Free Radical Scavenging Activity.” Journal of Agricultural
and Food Chemistry 54: 6969–6977. https://doi.org/10.1021/jf061090f.
Ferreira, D., J. P. Steynberg, J. F. W. Burger, and
B. C. B. Bezuidenhoudt. 1992. “Oxidation and Rearrangement Reactions
of Condensed Tannins.” In Plant Polyphenols: Synthesis, Properties,
Significance, edited by R. Hemingway, P. Laks and S. Branham,
349–384. Plenum. https://doi.org/10.1007/978-1-4615-3476-1_20.
Ferreira, F. P. S., S. R. Morais, M. T. F. Bara, et al. 2014. “Eugenia
calycina Cambess Extracts and Their Fractions: Their Antimicrobial
Activity and the Identification of Major Polar Compounds Using Elec-
trospray Ionization FT‐ICR Mass Spectrometry.” Journal of
Pharmaceutical and Biomedical Analysis 99: 89–96. https://doi.org/10.
1016/j.jpba.2014.07.003.
Flamini, R. 2003. “Mass Spectrometry in Grape and Wine Chemistry.
Part I: Polyphenols.” Mass Spectrometry Reviews 22: 218–250. https://
doi.org/10.1002/mas.10052.
Fracassetti, D., C. Costa, L. Moulay, and F. A. Tomás‐Barberán. 2013.
“Ellagic Acid Derivatives, Ellagitannins, Proanthocyanidins and Other
Phenolics, Vitamin C and Antioxidant Capacity of Two Powder Prod-
ucts From Camu‐Camu Fruit (Myrciaria dubia).” Food Chemistry 139:
578–588. https://doi.org/10.1016/j.foodchem.2013.01.121.
Franceschi, P., U. Vrhovsek, and G. Guella. 2011. “Ion Mobility Mass
Spectrometric Investigation of Ellagitannins and Their Non‐Covalent
Aggregates.” Rapid Communications in Mass Spectrometry 25: 827–833.
https://doi.org/10.1002/rcm.4932.
Freitas, V. A. P., Y. Glories, G. Bourgeois, and C. Vitry. 1998. “Char-
acterisation of Oligomeric and Polymeric Procyanidins From Grape
Seeds by Liquid Secondary Ion Mass Spectrometry.” Phytochemistry 49:
1435–1441. https://doi.org/10.1016/S0031-9422(98)00107-1.
Friedrich, W., A. Eberhardt, and R. Galensa. 2000. “Investigation of
Proanthocyanidins by HPLC With Electrospray Ionization Mass
21Mass Spectrometry Reviews, 2025
 10982787, 0, D
ow
nloaded from
 https://analyticalsciencejournals.onlinelibrary.wiley.com/doi/10.1002/mas.70013 by University of Turku, W
iley Online Library on [28/11/2025]. See the Terms and Conditions (https://onlinelibrary.wiley.com/terms-and-conditions) on W
iley Online Library for rules of use; OA articles are governed by the applicable Creative Commons License
Spectrometry.” European Food Research and Technology 211: 56–64.
https://doi.org/10.1007/s002170050589.
Fulcrand, H., S. Remy, J.‐M. Souquet, V. Cheynier, and M. Moutounet.
1999. “Study of Wine Tannin Oligomers by On‐Line Liquid Chroma-
tography Electrospray Ionization Mass Spectrometry.” Journal of
Agricultural and Food Chemistry 47: 1023–1028.
García, D. E., W. G. Glasser, A. Pizzi, S. P. Paczkowski, and M.‐P.
Laborie. 2016. “Modification of Condensed Tannins: From Polyphenol
Chemistry to Materials Engineering.” New Journal of Chemistry 40:
36–49. https://doi.org/10.1039/c5nj02131f.
García‐Estévez, I., M. T. Escribano‐Bailón, J. C. Rivas‐Gonzalo, and
C. Alcalde‐Eon. 2012. “Validation of a Mass Spectrometry Method to
Quantify Oak Ellagitannins in Wine Samples.” Journal of
Agricultural and Food Chemistry 60: 1373–1379. https://doi.org/10.
1021/jf203836a.
Gariboldi, E., D. Mascetti, G. Galli, P. Caballion, and E. Bosisio. 1998.
“LC‐UV‐Electrospray‐MS‐MS Mass Spectrometry Analysis of Plant
Constituents Inhibiting Xanthine Oxidase.” Pharmaceutical Research
15: 936–943. https://doi.org/10.1023/A:1011941002546.
Gasperotti, M., D. Masuero, F. Mattivi, and U. Vrhovsek. 2015. “Overall
Dietary Polyphenol Intake in a Bowl of Strawberries: The Influence of
Fragaria spp. in Nutritional Studies.” Journal of Functional Foods 18:
1057–1069. https://doi.org/10.1016/j.jff.2014.08.013.
Gaudin, E., M. Simon, J. Quijada, et al. 2016. “Efficacy of Sainfoin
(Onobrychis viciifolia) Pellets Against Multi Resistant Haemonchus
contortus and Interaction With Oral Ivermectin: Implications for On‐
Farm Control.” Veterinary Parasitology 227: 122–129. https://doi.org/10.
1016/j.vetpar.2016.08.002.
Ghahri, S., X. Chen, A. Pizzi, R. Hajihassani, and A. N. Papadopoulos.
2021. “Natural Tannins as New Cross‐Linking Materials for Soy‐Based
Adhesives.” Polymers 13: 595. https://doi.org/10.3390/polym13040595.
Giribaldi, J., M. Besson, L. Suc, H. Fulcrand, and L. Mouls. 2020. “The
Use of Extracted‐Ion Chromatograms to Quantify the Composition of
Condensed Tannin Subunits.” Rapid Communications in Mass
Spectrometry 34: e8619. https://doi.org/10.1002/rcm.8619.
Gong, Y., F. Fang, X. Zhang, et al. 2018. “B Type and Complex A/B Type
Epicatechin Trimers Isolated From Litchi pericarp Aqueous Extract
Show High Antioxidant and Anticancer Activity.” International Journal
of Molecular Sciences 19: 301. https://doi.org/10.3390/ijms19010301.
Gontijo, D. C., P. C. Gontijo, G. C. Brandão, et al. 2019. “Antioxidant
Study Indicative of Antibacterial and Antimutagenic Activities of an
Ellagitannin‐Rich Aqueous Extract From the Leaves of Miconia late-
crenata.” Journal of Ethnopharmacology 236: 114–123. https://doi.org/
10.1016/j.jep.2019.03.007.
Gross, E. M., A. Brune, and O. Walenciak. 2008. “Gut Ph, Redox
Conditions and Oxygen Levels in an Aquatic Caterpillar: Potential Ef-
fects on the Fate of Ingested Tannins.” Journal of Insect Physiology 54:
462–471. https://doi.org/10.1016/j.jinsphys.2007.11.005.
Gross, J. H. 2017. Mass Spectrometry. A Text Book (3rd ed.). Springer.
Gu, L., M. A. Kelm, J. F. Hammerstone, et al. 2003a. “Screening of
Foods Containing Proanthocyanidins and Their Structural Characteri-
zation Using LC‐MS/MS and Thiolytic Degradation.” Journal of
Agricultural and Food Chemistry 51: 7513–7521. https://doi.org/10.
1021/jf034815d.
Gu, L., M. A. Kelm, J. F. Hammerstone, et al. 2003b. “Liquid Chro-
matographic/Electrospray Ionization Mass Spectrometric Studies of
Proanthocyanidins in Foods.” Journal of Mass Spectrometry 38:
1272–1280. https://doi.org/10.1002/jms.541.
Guillarme, D., J. Schappler, S. Rudaz, and J.‐L. Veuthey. 2010. “Cou-
pling Ultra‐High‐Pressure Liquid Chromatography With Mass Spec-
trometry.” TrAC, Trends in Analytical Chemistry 29: 15–27. https://doi.
org/10.1016/j.trac.2009.09.008.
Gusev, A. I., W. R. Wilkinson, A. Proctor, and D. M. Hercules. 1995.
“Improvement of Signal Reproducibility and Matrix/Comatrix Effects in
MALDI Analysis.” Analytical Chemistry 67: 1034–1041. https://doi.org/
10.1021/ac00102a003.
Guyot, S., T. Doco, J.‐M. Souquet, M. Moutounet, and J.‐F. Drilleau.
1997. “Characterization of Highly Polymerized Procyanidins in Cider
Apple (Malus sylvestris Var. Kermerrien) Skin and Pulp.”
Phytochemistry 44: 351–357. https://doi.org/10.1016/S0031-9422(96)
00480-3.
Guyot, S., N. Marnet, D. Laraba, P. Sanoner, and J.‐F. Drilleau. 1998.
“Reversed‐Phase HPLC Following Thiolysis for Quantitative Estimation
and Characterization of the Four Main Classes of Phenolic Compounds
in Different Tissue Zones of a French Cider Apple Variety (Malus do-
mestica Var. Kermerrien).” Journal of Agricultural and Food Chemistry
46: 1698–1705. https://doi.org/10.1021/jf970832p.
Guyot, S., N. Marnet, P. Sanoner, and J.‐F. Drilleau. 2001. “Direct
Thiolysis on Crude Apple Materials for High‐Performance Liquid
Chromatography Characterization and Quantification of Polyphenols in
Cider Apple Tissues and Juices.” Methods in Enzymology 335: 57–70.
Guyot, S., J. Vercauteren, and V. Cheynier. 1996. “Colourless and Yel-
low Dimers Resulting From (+)‐Catechin Oxidative Coupling Catalysed
by Grape Polyphenoloxidase.” Phytochemistry 42: 1279–1288.
Hager, T. J., L. R. Howard, R. Liyanage, J. O. Lay, and R. L. Prior. 2008.
“Ellagitannin Composition of Blackberry as Determined by HPLC‐ESI‐
MS and MALDI‐TOF‐MS.” Journal of Agricultural and Food Chemistry
56: 661–669. https://doi.org/10.1021/jf071990b.
Hammerstone, J. F., S. A. Lazarus, A. E. Mitchell, R. Rucker, and
H. H. Schmitz. 1999. “Identification of Procyanidins in Cocoa
(Theobroma cacao) and Chocolate Using High‐Performance Liquid
Chromatography/Mass Spectrometry.” Journal of Agricultural and Food
Chemistry 47: 490–496.
Hatano, T., M. Hori, M. Kusuda, T. Ohyabu, H. Ito, and T. Yoshida.
2004. “Characterization of the Oxidation Products of (‐)‐
Epigallocatechin Gallate, a Bioactive Tea Polyphenol, on Incubation in
Neutral Solution.” Heterocycles 63: 1547–1554. https://doi.org/10.3987/
COM-04-10039.
Hayasaka, Y., E. J. Waters, V. Cheynier, M. J. Herderich, and S. Vidal.
2003. “Characterization of Proanthocyanidins in Grape Seeds Using
Electrospray Mass Spectrometry.” Rapid Communications in Mass
Spectrometry 17: 9–16. https://doi.org/10.1002/rcm.869.
He, J., A. R. F. Carvalho, N. Mateus, and V. De Freitas. 2010. “Spectral
Features and Stability of Oligomeric Pyranoanthocyanin‐Flavanol Pig-
ments Isolated From Red Wines.” Journal of Agricultural and Food
Chemistry 58: 9249–9258. https://doi.org/10.1021/jf102085e.
He, M., G. Peng, F. Xie, L. Hong, and Q. Cao. 2019. “Liquid
Chromatography–High‐Resolution Mass Spectrometry With ROI Strat-
egy for Non‐Targeted Analysis of the In Vivo/In Vitro Ingredients
Coming From Ligusticum chuanxiong Hort.” Chromatographia 82:
1069–1077. https://doi.org/10.1007/s10337-019-03740-x.
Hemingway, R. W. 1989a. “Structural Variation in Proanthocyanidins
and Their Derivatives.” In Chemistry and Significance of Condensed
Tannins, edited by R. W. Hemingway and J. J. Karchesy, 83–107.
Plenum Press.
Hemingway, R. W. 1989b. “Reactions at the Interflavanoid Bond of
Proanthocyanidins.” In Chemistry and Significance of Condensed Tan-
nins, edited by R. W. Hemingway and J. J. Karchesy, 265–283. Plenum
Press.
Hibi, Y., and E. Yanase. 2019. “Oxidation of Procyanidins With Various
Degrees of Condensation: Influence on the Color‐Deepening Phenom-
enon.” Journal of Agricultural and Food Chemistry 67: 4940–4946.
https://doi.org/10.1021/acs.jafc.9b02085.
Holčapek, M., R. Jirásko, and M. Lísa. 2010. “Basic Rules for the
Interpretation of Atmospheric Pressure Ionization Mass Spectra of
22 Mass Spectrometry Reviews, 2025
 10982787, 0, D
ow
nloaded from
 https://analyticalsciencejournals.onlinelibrary.wiley.com/doi/10.1002/mas.70013 by University of Turku, W
iley Online Library on [28/11/2025]. See the Terms and Conditions (https://onlinelibrary.wiley.com/terms-and-conditions) on W
iley Online Library for rules of use; OA articles are governed by the applicable Creative Commons License
Small Molecules.” Journal of Chromatography A 1217, no. Issue 25:
3908–3921. https://doi.org/10.1016/j.chroma.2010.02.049.
Hooi Poay, T., L. Sui Kiong, and C. Cheng Hock. 2011. “Characterisa-
tion of Galloylated Cyanogenic Glucosides and Hydrolysable Tannins
From Leaves of Phyllagathis rotundifolia by LC‐ESI‐MS/MS.”
Phytochemical Analysis 22: 516–525. https://doi.org/10.1002/pca.1312.
Hoste, H., F. Jackson, S. Athanasiadou, S. M. Thamsborg, and
S. O. Hoskin. 2006. “The Effects of Tannin‐Rich Plants on Parasitic
Nematodes in Ruminants.” Trends in Parasitology 22: 253–261. https://
doi.org/10.1016/j.pt.2006.04.004.
Imran, I. B., M. Karonen, J.‐P. Salminen, and M. T. Engström. 2021.
“Modification of Natural Proanthocyanidin Oligomers and Polymers via
Chemical Oxidation Under Alkaline Conditions.” ACS Omega 6:
4726–4739. https://doi.org/10.1021/acsomega.0c05515.
Johnson, K. S., and R. V. Barbehenn. 2000. “Oxygen Levels in the Gut
Lumens of Herbivorous Insects.” Journal of Insect Physiology 46:
897–903. https://doi.org/10.1016/S0022-1910(99)00196-1.
Jourdes, M., J. Michel, C. Saucier, S. Quideau, and P. L. Teissedre. 2011.
“Identification, Amounts, and Kinetics of Extraction of C‐Glucosidic
Ellagitannins During Wine Aging in Oak Barrels or in Stainless Steel
Tanks With Oak Chips.” Analytical and Bioanalytical Chemistry 401:
1531–1539. https://doi.org/10.1007/s00216-011-4949-8.
Kalili, K. M., and A. de Villiers. 2009. “Off‐Line Comprehensive
2‐Dimensional Hydrophilic Interaction × Reversed Phase Liquid
Chromatography Analysis of Procyanidins.” Journal of Chromatography
A 1216: 6274–6284. https://doi.org/10.1016/j.chroma.2009.06.071.
Kalili, K. M., and A. de Villiers. 2010. “Off‐Line Comprehensive Two‐
Dimensional Hydrophilic Interaction × Reversed Phase Liquid Chro-
matographic Analysis of Green Tea Phenolics.” Journal of Separation
Science 33: 853–863.
Karchesy, J. J., R. W. Hemingway, Y. L. Foo, E. Barofsky, and
D. F. Barofsky. 1986. “Sequencing Procyanidin Oligomers by Fast
Atom Bombardment Mass Spectrometry.” Analytical Chemistry 58:
2563–2567. https://doi.org/10.1021/ac00125a044.
Kårlund, A., J.‐P. Salminen, P. Koskinen, et al. 2014. “Polyphenols in
Strawberry (Fragaria × Ananassa) Leaves Induced by Plant Activators.”
Journal of Agricultural and Food Chemistry 62: 4592–4600. https://doi.
org/10.1021/jf405589f.
Karonen, M., J. R. Ahern, L. Legroux, et al. 2020. “Ellagitannins Inhibit
the Exsheathment Ofhaemonchus Contortusandtrichostrongylus Colu-
briformislarvae: The Efficiency Increases Together With the Molecular
Size.” Journal of Agricultural and Food Chemistry 68: 4176–4186.
Karonen, M., I. B. Imran, M. T. Engström, and J.‐P. Salminen. 2021.
“Characterization of Natural and Alkaline‐Oxidized Proanthocyanidins
in Plant Extracts by Ultrahigh‐Resolution UHPLC‐MS/MS.” Molecules
26: 1873.
Karonen, M., J. Liimatainen, and J. Sinkkonen. 2011. “Birch Inner Bark
Procyanidins Can Be Resolved With Enhanced Sensitivity by
Hydrophilic Interaction HPLC‐MS.” Journal of Separation Science 34:
3158–3165. https://doi.org/10.1002/jssc.201100569.
Karonen, M., J. Loponen, V. Ossipov, and K. Pihlaja. 2004. “Analysis of
Procyanidins in Pine Bark With Reversed‐Phase and Normal‐Phase
High‐Performance Liquid Chromatography‐Electrospray Ionization
Mass Spectrometry.” Analytica Chimica Acta 522: 105–112. https://doi.
org/10.1016/j.aca.2004.06.041.
Karonen, M., J. Parker, A. Agrawal, and J.‐P. Salminen. 2010. “First
Evidence of Hexameric and Heptameric Ellagitannins in Plants
Detected by Liquid Chromatography/Electrospray Ionisation Mass
Spectrometry.” Rapid Communications in Mass Spectrometry 24:
3151–3156. https://doi.org/10.1002/rcm.
Kelm, M. A., J. C. Johnson, R. J. Robbins, J. F. Hammerstone, and
H. H. Schmitz. 2006. “High‐Performance Liquid Chromatography
Separation and Purification of Cacao (Theobroma cacao L.) Procyani-
dins According to Degree of Polymerization Using a Diol Stationary
Phase.” Journal of Agricultural and Food Chemistry 54: 1571–1576.
https://doi.org/10.1021/jf0525941.
Kennedy, J. A., and G. P. Jones. 2001. “Analysis of Proanthocyanidin
Cleavage Products Following Acid‐Catalysis in the Presence of Excess
Phloroglucinol.” Journal of Agricultural and Food Chemistry 49:
1740–1746.
Kim, J., M. Pälijärvi, M. Karonen, and J.‐P. Salminen. 2018. “Oxidatively
Active Plant Phenolics Detected by UHPLC‐DAD‐MS After Enzymatic
and Alkaline Oxidation.” Journal of Chemical Ecology 44: 483–496.
https://doi.org/10.1007/s10886-018-0949-x.
Kim, J., M. Pälijärvi, M. Karonen, and J.‐P. Salminen. 2020. “Distribu-
tion of Enzymatic and Alkaline Oxidative Activities of Phenolic Com-
pounds in Plants.” Phytochemistry 179: 112501. https://doi.org/10.
1016/j.phytochem.2020.112501.
Kolodziej, H., O. Kayser, K. P. Latté, and A. Kiderlen. 1999. “En-
hancement of Antimicrobial Activity of tannins and Related Com-
pounds by Immune Modulatory Effects.” In Plant Polyphenols 2:
Chemistry, Biology, Ecology, edited by G. G. Gross, R. W. Hemingway,
and T. Yoshida, 575–594. Kluwer Academic/Plenum Publishers. https://
doi.org/10.1007/978-1-4615-4139-4.
Kondo, K., M. Kurihara, K. Fukuhara, et al. 2000. “Conversion of
Procyanidin B‐Type (Catechin Dimer) to A‐Type: Evidence for
Abstraction of C‐2 Hydrogen in Catechin During Radical Oxidation.”
Tetrahedron Letters 41: 485–488. https://doi.org/10.1016/S0040-4039(99)
02097-3.
Koupai‐Abyazani, M. R., J. McCallum, and B. A. Bohm. 1992. “Identi-
fication of the Constituent Flavanoid Units in Sainfoin Proanthocya-
nidins by Reversed‐Phase High‐Performance Liquid Chromatography.”
Journal of Chromatography A 594: 117–123. https://doi.org/10.1016/
0021-9673(92)80319-P.
Kpegba, K., A. Agbonon, A. G. Petrovic, et al. 2011. “Epiafzelechin
From the Root Bark of Cassia sieberiana: Detection by DART Mass
Spectrometry, Spectroscopic Characterization, and Antioxidant Prop-
erties.” Journal of Natural Products 74: 455–459. https://doi.org/10.
1021/np100090e.
Krueger, C. G., N. C. Dopke, P. M. Treichel, J. Folts, and J. D. Reed.
2000. “Matrix‐Assisted Laser Desorption/Ionization Time‐of‐Flight
Mass Spectrometry of Polygalloyl polyflavan‐3‐ols in Grape Seed Ex-
tract.” Journal of Agricultural and Food Chemistry 48: 1663–1667.
https://doi.org/10.1021/jf990534n.
Krueger, C. G., M. M. Vestling, and J. D. Reed. 2003. “Matrix‐Assisted
Laser Desorption/Ionization Time‐of‐Flight Mass Spectrometry of
Heteropolyflavan‐3‐ols and Glucosylated Heteropolyflavans In Sorghum
[Sorghum bicolor (L.) Moench].” Journal of Agricultural and Food
Chemistry 51: 538–543. https://doi.org/10.1021/jf020746b.
Kumar, S., A. Singh, V. Bajpai, B. Singh, and B. Kumar. 2017. “Devel-
opment of a UHPLC‐MS/MS Method for the Quantitation of Bioactive
Compounds in Phyllanthus Species and Its Herbal Formulations.”
Journal of Separation Science 40: 3422–3429. https://doi.org/10.1002/
jssc.201601361.
Laitila, J. E., and J.‐P. Salminen. 2020. “Relevance of the Concentrations
and Sizes of Oligomeric Red Wine Pigments to the Color Intensity of
Commercial Red Wines.” Journal of Agricultural and Food Chemistry
68: 3576–3584. https://doi.org/10.1021/acs.jafc.9b07941.
Laitila, J. E., J. Suvanto, and J.‐P. Salminen. 2019. “Liquid
Chromatography–Tandem Mass Spectrometry Reveals Detailed Chro-
matographic Fingerprints of Anthocyanins and Anthocyanin Adducts
in Red Wine.” Food Chemistry 294: 138–151. https://doi.org/10.1016/j.
foodchem.2019.02.136.
Laitila, J. E., P. T. Tähtinen, M. Karonen, and J.‐P. Salminen. 2023. “Red
Wine Inspired Chemistry: Hemisynthesis of Procyanidin Analogs and
23Mass Spectrometry Reviews, 2025
 10982787, 0, D
ow
nloaded from
 https://analyticalsciencejournals.onlinelibrary.wiley.com/doi/10.1002/mas.70013 by University of Turku, W
iley Online Library on [28/11/2025]. See the Terms and Conditions (https://onlinelibrary.wiley.com/terms-and-conditions) on W
iley Online Library for rules of use; OA articles are governed by the applicable Creative Commons License
Determination of Their Protein Precipitation Capacity, Octanol‐Water
Partition, and Stability in Phosphate‐Buffered Saline.” Journal of
Agricultural and Food Chemistry 71: 19832–19844. https://doi.org/10.
1021/acs.jafc.3c06467.
Laks, P. E., R. W. Hemingway, and A. H. Conner. 1987. “Condensed
Tannins. Base‐Catalysed Reactions of Polymeric Procyanidins With
Phloroglucinol: Intramolecular Rearrangements.” Journal of the
Chemical Society, Perkin Transactions.1: 1875–1881.
Lambert, M., E. Meudec, A. Verbaere, et al. 2015. “A High‐Throughput
UHPLC‐QQQ‐MS Method for Polyphenol Profiling in Rosé Wines.”
Molecules 20: 7890–7914. https://doi.org/10.3390/molecules20057890.
Lazarus, S. A., G. E. Adamson, J. F. Hammerstone, and H. H. Schmitz.
1999. “High‐Performance Liquid Chromatography/Mass Spectrometry
Analysis of Proanthocyanidins in Foods and Beverages.” Journal of
Agricultural and Food Chemistry 47: 3693–3701. https://doi.org/10.
1016/S0076-6879(01)35230-8.
Leppä, M. M., M. Karonen, P. Tähtinen, M. T. Engström, and J.‐P.
Salminen. 2018. “Isolation of Chemically Well‐Defined Semipreparative
Liquid Chromatography Fractions From Complex Mixtures of Proan-
thocyanidin Oligomers and Polymers.” Journal of Chromatography A
1576: 67–79. https://doi.org/10.1016/j.chroma.2018.09.034.
Liebler, D. C., and L. J. Zimmerman. 2013. “Targeted Quantitation of
Proteins by Mass Spectrometry.” Biochemistry 52: 3797–3806. https://
doi.org/10.1021/bi400110b.
Liimatainen, J., M. Karonen, and J. Sinkkonen. 2012. “Procyanidin
Xylosides From the Bark of Betula pendula.” Phytochemistry 76:
178–183. https://doi.org/10.1016/j.phytochem.2012.01.008.
Lin, L.‐Z., J. Sun, P. Chen, M. J. Monagas, and J. M. Harnly. 2014.
“UHPLC‐PDA‐ESI/HRMSn Profiling Method to Identify and Quantify
Oligomeric Proanthocyanidins in Plant Products.” Journal of
Agricultural and Food Chemistry 62: 9387–9400.
Lin, Y., W. Xu, W. Xu, et al. 2015. “Simultaneous Determination of 41
Components In Gualou Guizhi Granules by UPLC Coupled With Triple
Quadrupole Mass Spectrometry.” Analytical Methods 7: 8285–8296.
https://doi.org/10.1039/c5ay01336d.
Loo, J. A. 1997. “Studying Noncovalent Protein Complexes by Electro-
spray Ionization Mass Spectrometry.” Mass Spectrometry Reviews 16:
1–23. https://doi.org/10.1002/(SICI)1098-2787(1997)16:1<1::AID-
MAS1>3.0.CO;2-L.
López‐Andrés, P., G. Luciano, V. Vasta, et al. 2013. “Dietary Quebracho
Tannins Are Not Absorbed, but Increase the Antioxidant Capacity of
Liver and Plasma in Sheep.” British Journal of Nutrition 110: 632–639.
https://doi.org/10.1017/S0007114512005703.
Luo, F., Y. Fu, Y. Xiang, et al. 2014. “Identification and Quantification
of Gallotannins in Mango (Mangifera indica L.) Kernel and Peel and
Their Antiproliferative Activities.” Journal of Functional Foods 8:
282–291. https://doi.org/10.1016/j.jff.2014.03.030.
Maillard, M., P. Giampaoli, and M. Cuvelier. 1996. “Atmospheric
Pressure Chemical Ionization (APCI) Liquid Chromatography‐Mass
Spectrometry: Characterization of Natural Antioxidants.” Talanta 43:
339–347.
Maleknia, S. D., T. M. Vail, R. B. Cody, D. O. Sparkman, T. L. Bell, and
M. A. Adams. 2009. “Temperature‐Dependent Release of Volatile
Organic Compounds of Eucalypts by Direct Analysis in Real Time
(DART) Mass Spectrometry.” Rapid Communications in Mass
Spectrometry 23: 2241–2246. https://doi.org/10.1002/rcm.
Malik, G., A. Natangelo, J. Charris, et al. 2012. “Synthetic Studies
TowardC‐Glucosidic Ellagitannins: A Biomimetic Total Synthesis of
5‐O‐Desgalloylepipunicacortein A.” Chemistry – A European Journal 18:
9063–9074.
Mämmelä, P., H. Savolainen, L. Lindroos, J. Kangas, and T. Vartiainen.
2000. “Analysis of Oak Tannins by Liquid Chromatography‐
Electrospray Ionisation Mass Spectrometry.” Journal of
Chromatography A 891: 75–83. https://doi.org/10.1016/S0021-9673(00)
00624-5.
Mané, C., N. Sommerer, T. Yalcin, V. Cheynier, R. B. Cole, and
H. Fulcrand. 2007. “Assessment of the Molecular Weight Distribution of
Tannin Fractions Through MALDI‐TOF MS Analysis of Protein‐Tannin
Complexes.” Analytical Chemistry 79: 2239–2248. https://doi.org/10.
1021/ac061685+.
Manjón, E., I. García‐Estévez, and M. T. Escribano‐Bailón. 2024. “Pos-
sible Role of High‐Molecular‐Weight Salivary Proteins in Astringency
Development.” Foods 13: 862. https://doi.org/10.3390/foods13060862.
Margalef, M., Z. Pons, B. Muguerza, and A. Arola‐Arnal. 2014. “A Rapid
Method to Determine Colonic Microbial Metabolites Derived From
Grape Flavanols in Rat Plasma by Liquid Chromatography‐Tandem
Mass Spectrometry.” Journal of Agricultural and Food Chemistry 62:
7698–7706. https://doi.org/10.1021/jf5019752.
Matthews, S., I. Mila, A. Scalbert, et al. 1997. “Method for Estimation of
Proanthocyanidins Based on Their Acid Depolymerization in the
Presence of Nucleophiles.” Journal of Agricultural and Food Chemistry
45: 1195–1201.
Mau, M., A. M. de Almeida, A. V. Coelho, and K.‐H. Südekum. 2011.
“First Identification of Tannin‐Binding Proteins In Saliva of Papio
hamadryas Using MS/MS Mass Spectrometry.” American Journal of
Primatology 73: 896–902. https://doi.org/10.1002/ajp.20958.
Mazerolles, G., S. Preys, C. Bouchut, et al. 2010. “Combination of Sev-
eral Mass Spectrometry Ionization Modes: A Multiblock Analysis for a
Rapid Characterization of the Red Wine Polyphenolic Composition.”
Analytica Chimica Acta 678: 195–202. https://doi.org/10.1016/j.aca.
2010.07.034.
Mena, P., L. Calani, C. Dall'Asta, et al. 2012. “Rapid and Comprehensive
Evaluation of (Poly)Phenolic Compounds In Pomegranate (Punica
granatum L.) Juice by UHPLC‐MSn.” Molecules 17: 14821–14840.
https://doi.org/10.3390/molecules171214821.
Meyers, K. J., T. J. Swiecki, and A. E. Mitchell. 2006. “Understanding
the Native Californian Diet: Identification of Condensed and Hydro-
lyzable Tannins in Tanoak Acorns (Lithocarpus densiflorus).” Journal
of Agricultural and Food Chemistry 54: 7686–7691. https://doi.org/10.
1021/jf061264t.
Moilanen, J., and J.‐P. Salminen. 2008. “Ecologically Neglected Tannins
and Their Biologically Relevant Activity: Chemical Structures of Plant
Ellagitannins Reveal Their In Vitro Oxidative Activity at High pH.”
Chemoecology 18: 73–83. https://doi.org/10.1007/s00049-007-0395-7.
Moilanen, J., J. Sinkkonen, and J.‐P. Salminen. 2013. “Characterization
of Bioactive Plant Ellagitannins by Chromatographic, Spectroscopic and
Mass Spectrometric Methods.” Chemoecology 23: 165–179. https://doi.
org/10.1007/s00049-013-0132-3.
Monagas, M., J. E. Quintanilla‐López, C. Gómez‐Cordovés,
B. Bartolomé, and R. Lebrón‐Aguilar. 2010. “MALDI‐TOF MS Analysis
of Plant Proanthocyanidins.” Journal of Pharmaceutical and Biomedical
Analysis 51: 358–372. https://doi.org/10.1016/j.jpba.2009.03.035.
Mouls, L., and H. Fulcrand. 2012. “Uplc‐Esi‐Ms Study of the Oxidation
Markers Released From Tannin Depolymerization: Toward a Better
Characterization of the Tannin Evolution Over Food and Beverage
Processing.” Journal of Mass Spectrometry 47: 1450–1457. https://doi.
org/10.1002/jms.3098.
Mouls, L., and H. Fulcrand. 2015. “Identification of New Oxidation
Markers of Grape‐Condensed Tannins by UPLC‐MS Analysis After
Chemical Depolymerization.” Tetrahedron 71: 3012–3019. https://doi.
org/10.1016/j.tet.2015.01.038.
Mouls, L., J. P. Mazauric, N. Sommerer, H. Fulcrand, and G. Mazerolles.
2011. “Comprehensive Study of Condensed Tannins by ESI Mass
Spectrometry: Average Degree of Polymerisation and Polymer
24 Mass Spectrometry Reviews, 2025
 10982787, 0, D
ow
nloaded from
 https://analyticalsciencejournals.onlinelibrary.wiley.com/doi/10.1002/mas.70013 by University of Turku, W
iley Online Library on [28/11/2025]. See the Terms and Conditions (https://onlinelibrary.wiley.com/terms-and-conditions) on W
iley Online Library for rules of use; OA articles are governed by the applicable Creative Commons License
Distribution Determination From Mass Spectra.” Analytical and
Bioanalytical Chemistry 400: 613–623. https://doi.org/10.1007/s00216-
011-4751-7.
Mullen, W., T. Yokota, M. E. J. Lean, and A. Crozier. 2003. “Analysis of
Ellagitannins and Conjugates of Ellagic Acid and Quercetin in Rasp-
berry Fruits by LC‐MSn.” Phytochemistry 64: 617–624. https://doi.org/
10.1016/S0031-9422(03)00281-4.
Navarro‐Hoyos, M., R. Lebrón‐Aguilar, J. E. Quintanilla‐López, et al.
2017. “Proanthocyanidin Characterization and Bioactivity of Extracts
From Different Parts of Uncaria tomentosa L. (Cat's Claw).”
Antioxidants 6: 12.
Nguyen, G. T. H., J. L. Bennett, S. Liu, et al. 2021. “Multiplexed
Screening of Thousands of Natural Products for Protein–Ligand Binding
in Native Mass Spectrometry.” Journal of the American Chemical Society
143: 21379–21387. https://doi.org/10.1021/jacs.1c10408.
Niu, S., J. N. Rabuck, and B. T. Ruotolo. 2013. “Ion Mobility‐Mass
Spectrometry of Intact Protein‐Ligand Complexes for Pharmaceutical
Drug Discovery and Development.” Current Opinion in Chemical
Biology 17: 809–817. https://doi.org/10.1016/j.cbpa.2013.06.019.
Nonaka, G., O. Kawahara, and I. Nishioka. 1983. “Tannins and Related
Compounds. XV. A New Class of Dimeric Flavan‐3‐ol Gallates, Thea-
sinensins A and B, and Proanthocyanidin Gallates From Green Tea
Leaf.” Chemical and Pharmaceutical Bulletin 31: 3906–3914.
Ontañón, I., D. Sánchez, V. Sáez, F. Mattivi, V. Ferreira, and
P. Arapitsas. 2020. “Liquid Chromatography‐Mass Spectrometry‐Based
Metabolomics for Understanding the Compositional Changes Induced
by Oxidative or Anoxic Storage of Red Wines.” Journal of Agricultural
and Food Chemistry 68: 13367–13379. https://doi.org/10.1021/acs.jafc.
0c04118.
Ortega, N., M.‐P. Romero, A. Macià, et al. 2010. “Comparative Study of
UPLC‐MS/MS and HPLC‐MS/MS to Determine Procyanidins and Al-
kaloids in Cocoa Samples.” Journal of Food Composition and Analysis
23: 298–305. https://doi.org/10.1016/j.jfca.2009.10.005.
Palanisamy, U. D., L. T. Ling, T. Manaharan, and D. Appleton. 2011.
“Rapid Isolation of Geraniin From Nephelium lappaceum Rind Waste
and Its Anti‐Hyperglycemic Activity.” Food Chemistry 127: 21–27.
https://doi.org/10.1016/j.foodchem.2010.12.070.
Perez‐Gregorio, M. R., N. Mateus, and V. de Freitas. 2014. “New Pro-
cyanidin B3‐Human Salivary Protein Complexes by Mass Spectrometry.
Effect of Salivary Protein Profile, Tannin Concentration, and Time
Stability.” Journal of Agricultural and Food Chemistry 62: 10038–10045.
https://doi.org/10.1021/jf5033284.
Pérez‐Jiménez, J., and J. L. Torres. 2012. “Analysis of Proanthocyani-
dins in Almond Blanch Water by HPLC‐ESI‐QQQ‐MS/MS and MALDI‐
TOF/TOF MS.” Food Research International 49: 798–806. https://doi.
org/10.1016/j.foodres.2012.09.005.
Pérez‐Magariño, S., I. Revilla, M. L. González‐Sanjosé, and S. Beltrán.
1999. “Various Applications of Liquid Chromatography‐Mass Spec-
trometry to the Analysis of Phenolic Compounds.” Journal of
Chromatography A 847: 75–81. https://doi.org/10.1016/S0021-9673(99)
00255-1.
Petit, E., R. Jacquet, L. Pouységu, D. Deffieux, and S. Quideau. 2019.
“Reactivity of Wine Polyphenols under Oxidation Conditions: Hemi-
synthesis of Adducts Between Grape Catechins or Oak Ellagitannins
and Odoriferous Thiols.” Tetrahedron 75: 551–560. https://doi.org/10.
1016/j.tet.2018.11.071.
Petit, E., D. Lefeuvre, R. Jacquet, L. Pouységu, D. Deffieux, and
S. Quideau. 2013. “Remarkable Biomimetic Chemoselective Aerobic
Oxidation of Flavano‐Ellagitannins Found in Oak‐Aged Wine.”
Angewandte Chemie International Edition 52: 11530–11533.
Pfundstein, B., S. K. El Desouky, W. E. Hull, R. Haubner, G. Erben, and
R. W. Owen. 2010. “Polyphenolic Compounds in the Fruits of Egyptian
Medicinal Plants (Terminalia bellerica, Terminalia chebulaand
Terminalia horrida): Characterization, Quantitation and Determination
of Antioxidant Capacities.” Phytochemistry 71: 1132–1148. https://doi.
org/10.1016/j.phytochem.2010.03.018.
Porter, L. J. 1989a. “Tannins.” In Methods in Plant Biochemistry, edited
by P. M. Dey and J. B. Harborne, 1, 389–419. Academic Press.
Porter, L. J. 1989b. “Condensed tannins.” In Natural Products of Woody
Plants I, edited by J. W. Rowe, 651–690. Springer‐Verlag.
Pouységu, L., D. Deffieux, G. Malik, A. Natangelo, and S. Quideau.
2011. “Synthesis of Ellagitannin Natural Products.” Natural Product
Reports 28: 853–874. https://doi.org/10.1039/c0np00058b.
Puljula, E., G. Walton, M. J. Woodward, and M. Karonen. 2020. “An-
timicrobial Activities of Ellagitannins Against Clostridiales perfringens,
Escherichia coli, Lactobacillus plantarum and Staphylococcus aureus.”
Molecules 25: 3714. https://doi.org/10.3390/molecules25163714.
Quideau, S., D. Deffieux, C. Douat‐Casassus, and L. Pouységu. 2011.
“Plant Polyphenols: Chemical Properties, Biological Activities, and
Synthesis.” Angewandte Chemie International Edition 50: 586–621.
https://doi.org/10.1002/anie.201000044.
Quideau, S., M. Jourdes, D. Lefeuvre, et al. 2005. “The Chemistry of
Wine Polyphenolic C‐Glycosidic Ellagitannins Targeting Human
Topoisomerase Ii.” Chemistry – A European Journal 11: 6503–6513.
Quideau, S., M. Jourdes, C. Saucier, Y. Glories, P. Pardon, and
C. Baudry. 2003. “Dna Topoisomerase Inhibitor Acutissimin A and
Other Flavano‐Ellagitannins in Red Wine.” Angewandte Chemie
International Edition 42: 6012–6014.
Quijada, J., C. Fryganas, H. M. Ropiak, A. Ramsay, I. Mueller‐Harvey,
and H. Hoste. 2015. “Anthelmintic Activities Against Haemonchus
contortus or Trichostrongylus colubriformis From Small Ruminants
Are Influenced by Structural Features of Condensed Tannins.” Journal
of Agricultural and Food Chemistry 63: 6346–6354. https://doi.org/10.
1021/acs.jafc.5b00831.
Rasines‐Perea, Z., R. Jacquet, M. Jourdes, S. Quideau, and P. L. Teissedre.
2019. “Ellagitannins and Flavano‐Ellagitannins: Red Wines Tendency in
Different Areas, Barrel Origin and Ageing Time in Barrel and Bottle.”
Biomolecules 9: 316. https://doi.org/10.3390/biom9080316.
Rauha, J. P., J. L. Wolfender, J. P. Salminen, K. Pihlaja, K. Hostettmann,
and H. Vuorela. 2001. “Characterization of the Polyphenolic Compo-
sition of Purple Loosestrife (Lythrum salicaria).” Zeitschrift fur
Naturforschung ‐ Section C Journal of Biosciences 56: 13–20. https://
doi.org/10.1515/znc-2001-1-203.
Reddy, M., S. Gupta, M. Jacob, S. Khan, and D. Ferreira. 2007. “Anti-
oxidant, Antimalarial and Antimicrobial Activities of Tannin‐Rich
Fractions, Ellagitannins and Phenolic Acids From Punica granatum L.”
Planta Medica 73: 461–467. https://doi.org/10.1055/s-2007-967167.
Reed, J. D., C. G. Krueger, M. M. Vestling, et al. 2005. “Maldi‐Tof Mass
Spectrometry of Oligomeric Food Polyphenols.” International Food
Research Journal 20: 2023–2034. https://doi.org/10.1016/j.phytochem.
2005.05.015.
Regazzoni, L., E. Arlandini, D. Garzon, N. A. Santagati, G. Beretta, and
R. Maffei Facino. 2013. “A Rapid Profiling of Gallotannins and Flavo-
noids of the Aqueous Extract of Rhus coriaria L. by Flow Injection
Analysis With High‐Resolution Mass Spectrometry Assisted With Da-
tabase Searching.” Journal of Pharmaceutical and Biomedical Analysis
72: 202–207. https://doi.org/10.1016/j.jpba.2012.08.017.
Regueiro, J., C. Sánchez‐González, A. Vallverdú‐Queralt, J. Simal‐
Gándara, R. Lamuela‐Raventós, and M. Izquierdo‐Pulido. 2014.
“Comprehensive Identification of Walnut Polyphenols by Liquid
Chromatography Coupled to Linear Ion Trap‐Orbitrap Mass Spec-
trometry.” Food Chemistry 152: 340–348. https://doi.org/10.1016/j.
foodchem.2013.11.158.
Robbins, K. S., Y. Ma, M. L. Wells, P. Greenspan, and R. B. Pegg. 2014.
“Separation and Characterization of Phenolic Compounds From U.S.
25Mass Spectrometry Reviews, 2025
 10982787, 0, D
ow
nloaded from
 https://analyticalsciencejournals.onlinelibrary.wiley.com/doi/10.1002/mas.70013 by University of Turku, W
iley Online Library on [28/11/2025]. See the Terms and Conditions (https://onlinelibrary.wiley.com/terms-and-conditions) on W
iley Online Library for rules of use; OA articles are governed by the applicable Creative Commons License
Pecans by Liquid Chromatography‐Tandem Mass Spectrometry.”
Journal of Agricultural and Food Chemistry 62: 4332–4341. https://doi.
org/10.1016/j.chroma.2014.06.027.
Roux, E. L., T. Doco, P. Sarni‐Manchado, Y. Lozano, and V. Cheynier.
1998. “A‐Type Proanthocyanidins From Pericarp of Litchi chinensis.”
Phytochemistry 48: 1251–1258.
Rue, E. A., M. D. Rush, and R. B. van Breemen. 2018. “Procyanidins: A
Comprehensive Review Encompassing Structure Elucidation via Mass
Spectrometry.” Phytochemistry Reviews 17: 1–16. https://doi.org/10.
1007/s11101-017-9507-3.
Saha, S., and I. B. Imran. 2020. “Isolation, Detection, and Quantification
of Hydrolyzable Tannins of the Biosynthetic Pathway by Liquid Chro-
matography Coupled With Tandem Mass Spectrometry.” Rapid
Communications in Mass Spectrometry 35: e9005. https://doi.org/10.
1002/rcm.9005.
Salminen, J.‐P. 2018. “Two‐Dimensional Tannin Fingerprints by Liquid
Chromatography Tandem Mass Spectrometry Offer a New Dimension
to Plant Tannin Analyses and Help to Visualize the Tannin Diversity n
Plants.” Journal of Agricultural and Food Chemistry 66: 9162–9171.
https://doi.org/10.1021/acs.jafc.8b02115.
Salminen, J.‐P., and M. Karonen. 2011. “Chemical Ecology of Tannins
and Other Phenolics: We Need a Change in Approach.” Functional
Ecology 25: 325–338. https://doi.org/10.1111/j.1365-2435.2010.01826.x.
Salminen, J.‐P., M. Karonen, K. Lempa, et al. 2005. “Characterisation of
Proanthocyanidin Aglycones and Glycosides From Rose Hips by High‐
Performance Liquid Chromatography‐Mass Spectrometry, and Their
Rapid Quantification Together With Vitamin C.” Journal of
Chromatography A 1077: 170–180. https://doi.org/10.1016/j.chroma.
2005.04.073.
Salminen, J.‐P., M. Karonen, and J. Sinkkonen. 2011. “Chemical Ecol-
ogy of Tannins: Recent Developments in Tannin Chemistry Reveal New
Structures and Structure‐Activity Patterns.” Chemistry – A European
Journal 17: 2806–2816. https://doi.org/10.1002/chem.201002662.
Salminen, J. P., and K. Lempa. 2002. “Effects of Hydrolysable Tannins
on a Herbivorous Insect: Fate of Individual Tannins in Insect Digestive
Tract.” Chemoecology 12: 203–211. https://doi.org/10.1007/PL00012670.
Salminen, J.‐P., V. Ossipov, J. Loponen, E. Haukioja, and K. Pihlaja.
1999. “Characterisation of Hydrolysable Tannins From Leaves of Betula
pubescens by High‐Performance Liquid Chromatography‐Mass Spec-
trometry.” Journal of Chromatography A 864: 283–291. https://doi.org/
10.1016/S0021-9673(99)01036-5.
Sargent, M., 2013. “Guide to Achieving Reliable Quantitative LC‐MS
Measurements, RSC Analytical Methods Committee.”
Sarni‐Manchado, P., and V. Cheynier. 2002. “Study of Non‐Covalent
Complexation Between Catechin Derivatives and Peptides by Electro-
spray Ionization Mass Spectrometry.” Journal of Mass Spectrometry 37:
609–616. https://doi.org/10.1002/jms.321.
Saucier, C., M. Jourdes, Y. Glories, and S. Quideau. 2006. “Extraction,
Detection, and Quantification of Flavano‐Ellagitannins and Ethylves-
calagin in a Bordeaux Red Wine Aged in Oak Barrels.” Journal of
Agricultural and Food Chemistry 54: 7349–7354. https://doi.org/10.
1021/jf061724i.
Scalbert, A. 1991. “Antimicrobial Properties of Tannins.”
Phytochemistry 30: 3875–3883. https://doi.org/10.1016/0031-9422(91)
83426-L.
Seeram, N., R. Lee, M. Hardy, and D. Heber. 2005. “Rapid Large Scale
Purification of Ellagitannins From Pomegranate Husk, a By‐Product of
the Commercial Juice Industry.” Separation and Purification
Technology 41: 49–55. https://doi.org/10.1016/j.seppur.2004.04.003.
Shraberg, J., S. W. Rick, N. Rannulu, and R. B. Cole. 2015. “A Study of
Procyanidin Binding to Histatin 5 Using Electrospray Ionization Tan-
dem Mass Spectrometry (ESI‐MS/MS) and Molecular Simulations.”
Physical Chemistry Chemical Physics 17: 12247–12258. https://doi.org/
10.1039/c4cp05586a.
Simon, C., K. Barathieu, M. Laguerre, et al. 2003. “Three‐Dimensional
Structure and Dynamics of Wine Tannin‐Saliva Protein Complexes. A
Multitechnique Approach.” Biochemistry 42: 10385–10395. https://doi.
org/10.1021/bi034354p.
Singh, A., V. Bajpai, S. Kumar, K. R. Sharma, and B. Kumara. 2016. “Pro-
filing of Gallic and Ellagic Acid Derivatives in Different Plant Parts of Ter-
minalia arjuna by HPLC‐ESI‐QTOF‐MS/MS.” Natural Product
Communications 11: 239–244. https://doi.org/10.1177/1934578x1601100227.
Sobeh, M., S. Rezq, O. M. Sabry, et al. 2019. “Albizia Anthelmintica:
HPLC‐MS/MS Profiling and In Vivo Anti‐Inflammatory, Pain Killing
and Antipyretic Activities of Its Leaf Extract.” Biomedicine &
Pharmacotherapy 115: 108882. https://doi.org/10.1016/j.biopha.2019.
108882.
Soong, Y. Y., and P. J. Barlow. 2005. “Isolation and Structure Elucida-
tion of Phenolic Compounds From Longan (Dimocarpus longan Lour.)
Seed by High‐Performance Liquid Chromatography‐ Electrospray
Ionization Mass Spectrometry.” Journal of Chromatography A 1085:
270–277. https://doi.org/10.1016/j.chroma.2005.06.042.
Soorkia, S., C. Jouvet, and G. Grégoire. 2020. “UV Photoinduced
Dynamics of Conformer‐Resolved Aromatic Peptides.” Chemical
Reviews 120: 3296–3327. https://doi.org/10.1021/acs.chemrev.9b00316.
Stark, T., N. Wollmann, K. Wenker, S. Lösch, A. Glabasnia, and
T. Hofmann. 2010. “Matrix‐Calibrated LC‐MS/MS Quantitation and
Sensory Evaluation of Oak Ellagitannins and Their Transformation
Products in Red Wines.” Journal of Agricultural and Food Chemistry 58:
6360–6369. https://doi.org/10.1021/jf100884y.
Sui, Y., Y. Zheng, X. Li, S. Li, B. Xie, and Z. Sun. 2016. “Characterization
and Preparation of Oligomeric Procyanidins From Litchi chinensis
Pericarp.” Fitoterapia 112: 168–174. https://doi.org/10.1016/j.fitote.
2016.06.001.
Sylla, T., L. Pouységu, G. Da Costa, D. Deffieux, J. P. Monti, and
S. Quideau. 2015. “Gallotannins and Tannic Acid: First Chemical
Syntheses and In Vitro Inhibitory Activity on Alzheimer's Amyloid β‐
Peptide Aggregation.” Angewandte Chemie International Edition 54:
8217–8221. https://doi.org/10.1002/anie.201411606.
Tanaka, T., G. Nonaka, and I. Nishioka. 1990. “Tannins and Related
Compounds. C. Reaction of Dehydrohexahydroxydiphenic Acid Esters
With Bases, and Its Application to the Structure Determination of
Pomegranate Tannins, Granatins A and B.” Chemical and
Pharmaceutical Bulletin 38: 2424–2428.
Tang, C., T. Guo, Z. Zhang, P. Yang, and H. Song. 2021. “Rapid Visu-
alized Characterization of Phenolic Taste Compounds in Tea Extract by
High‐Performance Thin‐Layer Chromatography Coupled to Desorption
Electrospray Ionization Mass Spectrometry.” Food Chemistry 355:
129555. https://doi.org/10.1016/j.foodchem.2021.129555.
Taylor, V. F., R. E. March, H. P. Longerich, and C. J. Stadey. 2005. “A
Mass Spectrometric Study of Glucose, Sucrose, and Fructose Using an
Inductively Coupled Plasma and Electrospray Ionization.” International
Journal of Mass Spectrometry 243: 71–84. https://doi.org/10.1016/j.ijms.
2005.01.001.
Thompson, R. S., D. Jacques, E. Haslam, and R. J. N. Tanner. 1972.
“Plant Proanthocyanidins. Part I. Introduction; the Isolation, Structure
and Distribution in Nature of Plant Procyanidins.” Journal of the
Chemical Society, Perkin Transactions 1 11: 1387–1399.
Tsakalof, A., A. A. Sysoev, K. V. Vyatkina, et al. 2024. “Current Role and
Potential of Triple Quadrupole Mass Spectrometry in Biomedical
Research and Clinical Applications.” Molecules 29: 5808. https://doi.
org/10.3390/molecules29235808.
Tuominen, A., and J. P. Salminen. 2017. “Hydrolyzable Tannins, Fla-
vonol Glycosides, and Phenolic Acids Show Seasonal and Ontogenic
26 Mass Spectrometry Reviews, 2025
 10982787, 0, D
ow
nloaded from
 https://analyticalsciencejournals.onlinelibrary.wiley.com/doi/10.1002/mas.70013 by University of Turku, W
iley Online Library on [28/11/2025]. See the Terms and Conditions (https://onlinelibrary.wiley.com/terms-and-conditions) on W
iley Online Library for rules of use; OA articles are governed by the applicable Creative Commons License
Variation in Geranium sylvaticum.” Journal of Agricultural and Food
Chemistry 65: 6387–6403. https://doi.org/10.1021/acs.jafc.7b00918.
Tuominen, A., and T. Sundman. 2013. “Stability and Oxidation Prod-
ucts of Hydrolysable Tannins in Basic Conditions Detected by HPLC/
DAD‐ESI/QTOF/MS.” Phytochemical Analysis 24: 424–435. https://doi.
org/10.1002/pca.2456.
Tuominen, A., E. Toivonen, P. Mutikainen, and J.‐P. Salminen. 2013.
“Defensive Strategies In Geranium sylvaticum. Part 1: Organ‐Specific
Distribution of Water‐Soluble Tannins, Flavonoids and Phenolic Acids.”
Phytochemistry 95: 394–407. https://doi.org/10.1016/j.phytochem.2013.
05.013.
Ventura, G., C. D. Calvano, G. Bianco, A. Di Capua, I. Losito, and
T. R. I. Cataldi. 2024. “Discovering the Ellagitannin Landscape of Dried
Walnut Shells by Liquid Chromatography With Tandem Mass Spec-
trometry.” Journal of Agricultural and Food Chemistry 72: 19051–19060.
Vergé, S., T. Richard, S. Moreau, et al. 2002. “First Observation of Non‐
Covalent Complexes for a Tannin‐Protein Interaction Model Investi-
gated by Electrospray Ionisation Mass Spectroscopy.” Tetrahedron
Letters 43: 2363–2366. https://doi.org/10.1016/S0040-4039(02)00255-1.
Vihakas, M., I. Gómez, M. Karonen, P. Tähtinen, I. Sääksjärvi, and J.‐P.
Salminen. 2015. “Phenolic Compounds and Their Fates in Tropical
Lepidopteran Larvae: Modifications in Alkaline Conditions.” Journal of
Chemical Ecology 41: 822–836. https://doi.org/10.1007/s10886-
015-0620-8.
Vihakas, M., M. Pälijärvi, M. Karonen, H. Roininen, and J.‐P. Salminen.
2014. “Rapid Estimation of the Oxidative Activities of Individual Phe-
nolics in Crude Plant Extracts.” Phytochemistry 103: 76–84. https://doi.
org/10.1016/j.phytochem.2014.02.019.
Vinci, G., A. Piccolo, and M. Bridoux. 2022. “Complementary ESI and
APPI High Resolution Mass Spectrometry Unravel the Molecular
Complexity of a Soil Humeome.” Analytica Chimica Acta 1194: 339398.
https://doi.org/10.1016/j.aca.2021.339398.
Vivas, N., G. Bourgeois, C. Vitry, Y. Glories, and V. Freitas. 1996.
“Determination of the Composition of Commercial Tannin Extracts by
Liquid Secondary Ion Mass Spectrometry (LSIMS).” Journal of the
Science of Food and Agriculture 72: 309–317. https://doi.org/10.1002/
(SICI)1097-0010(199611)72:3<309::AID-JSFA658>3.0.CO;2-U.
Weber, H. A., A. E. Hodges, J. R. Guthrie, et al. 2007. “Comparison of
Proanthocyanidins in Commercial Antioxidants: Grape Seed and Pine
Bark Extracts.” Journal of Agricultural and Food Chemistry 55: 148–156.
https://doi.org/10.1021/jf063150n.
Wu, L., K. Qi, C. Liu, Y. Hu, M. Xu, and Y. Pan. 2022. “Enhanced
Coverage and Sensitivity of Imprint DESI Mass Spectrometry Imaging
for Plant Leaf Metabolites by Post‐Photoionization.” Analytical
Chemistry 94: 15108–15116. https://doi.org/10.1021/acs.analchem.
2c03329.
Yang, B., M. Kortesniemi, P. Liu, M. Karonen, and J.‐P. Salminen. 2012.
“Analysis of Hydrolyzable Tannins and Other Phenolic Compounds in
Emblic Leafflower (Phyllanthus emblica L.) Fruits by High‐
Performance Liquid Chromatography‐Electrospray Ionization Mass
Spectrometry.” Journal of Agricultural and Food Chemistry 60:
8672–8683. https://doi.org/10.1021/jf302925v.
Yang, Y., and M. Chien. 2000. “Characterization of Grape Procyanidins
Using High‐Performance Liquid Chromatography/Mass Spectrometry
and Matrix‐Assisted Laser Desorption/Ionization Time‐of‐Flight
Mass Spectrometry.” Journal of Agricultural and Food Chemistry 48:
3990–3996. https://doi.org/10.1021/jf000316q.
Yoshida, T., K. Tanaka, X. M. Chen, and T. Okuda. 1991. “Tannins
From Hippophae rhamnoides.” Phytochemistry 30: 663–666.
Zhao, J., J. Wang, Y. Chen, and R. Agarwal. 1999. “Anti‐Tumor‐
Promoting Activity of a Polyphenolic Fraction Isolated From Grape
Seeds in the Mouse Skin Two‐Stage Initiation‐Promotion Protocol and
Identification of Procyanidin B5‐3ʹ‐Gallate as the Most Effective
Antioxidant Constituent.” Carcinogenesis 20: 1737–1745. https://doi.
org/10.1093/carcin/20.9.1737.
Zywicki, B., T. Reemtsma, and M. Jekel. 2002. “Analysis of Commercial
Vegetable Tanning Agents by Reversed‐Phase Liquid Chromatography‐
Electrospray Ionization‐Tandem Mass Spectrometry and Its Application
to Wastewater.” Journal of Chromatography A 970: 191–200. https://doi.
org/10.1016/S0021-9673(02)00883-X.
27Mass Spectrometry Reviews, 2025
 10982787, 0, D
ow
nloaded from
 https://analyticalsciencejournals.onlinelibrary.wiley.com/doi/10.1002/mas.70013 by University of Turku, W
iley Online Library on [28/11/2025]. See the Terms and Conditions (https://onlinelibrary.wiley.com/terms-and-conditions) on W
iley Online Library for rules of use; OA articles are governed by the applicable Creative Commons License