Soluble xyloglucan generates bigger bacterial community shifts than pectic polymers during in vitro fecal fermentation

Carbohydrate Polymers 206 (2019) 389–395

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Carbohydrate Polymers
journal homepage: www.elsevier.com/locate/carbpol

Soluble xyloglucan generates bigger bacterial community shifts than pectic
polymers during in vitro fecal fermentation

T

Thaisa Moro Cantu-Junglesa,b, Geórgia Erdmman do Nascimentoa, Xiaowei Zhangb,
⁎
⁎⁎
Marcello Iacominia, Lucimara M.C. Cordeiroa, , Bruce R. Hamakerb,
a
b

Departamento de Bioquímica e Biologia Molecular, Universidade Federal do Paraná, Curitiba, Brazil
Whistler Center for Carbohydrate Research and Department of Food Science, Purdue University, West Lafayette, USA

A R T I C LE I N FO

A B S T R A C T

Keywords:
In vitro fecal fermentation
Pectin
Xyloglucan
Short chain fatty acids

Prebiotics

Xyloglucans and pectic polymers can be obtained from a variety of plants ubiquitous in the human diet, however, their fermentability in the colon and consequent nutritional benefits are poorly understood. Here, we
evaluated metabolite profiles and bacterial shifts during in vitro fecal fermentations of two isolated pectic
polymers and a xyloglucan. Depending on their chemical structure, pectic polymers were more acetogenic or
propiogenic. Xyloglucan fermentation also resulted in elevated propionate if compared to FOS. Bacteroides
plebeius, B. uniformis, Parabacteroides distasonis and bacterial groups such as Blautia, Lachnospira, Clostridiales and
Lachnospiraceae, presented distinct abundances on each dietary fiber ferment. PCA and heat map analysis
showed that major microbiota shifts occurred during xyloglucan fermentation, but not pectin fermentation.
These data suggest that uncommon carbohydrate structures (i.e. isolated, soluble xyloglucan) in the diet hold the
potential to generate larger shifts in microbiota communities than commonly consumed fibers (i.e. pectins).

1. Introduction
Dietary fiber is the part of the plant material resistant to enzymatic
digestion in the small intestine and, thus, arrives intact in the colon
where may be fermented by resident bacteria (Codex Alimentarius
Commission, 2009). Saccharification and fermentation of some dietary
fibers in the large intestine result in the generation of beneficial metabolites such as short chain fatty acids (mainly acetate, butyrate and
propionate) and/or lead to bacterial shifts that are of biological significance to the human health (den Besten et al., 2013; Holscher, 2017).
It was proposed that depending on specific structural features of a
dietary fiber, a distinct fermentation profile would be observed
(Hamaker & Tuncil, 2014; Holscher, 2017). Dietary fiber composition
in higher plants varies among different plant types and cell types within
individual plants. In general, most dicotyledonous and non-graminaceous monocotyledons species possess xyloglucans as the predominant
hemicellulose. These species are also relatively rich in pectins, which
make up ∼35% of cell wall dry mass (Carpita & Gibeaut, 1993; OchoaVillarreal, Aispuro-Hernández, Vargas-Arispuro, & Ángel MartínezTéllez, 2012; Willats, Knox, & Mikkelsen, 2006).
Xyloglucans are strongly associated with cellulose and thus

⁎

entrapped in the insoluble cell wall matrix. When isolated, however,
these polymers became water soluble. This soluble form is hardly found
in the intact plant cell wall and consequently rarely consumed in a
regular diet. Xyloglucans have a (1→4)-linked β-D-Glcp backbone with
α-D-Xylp substituents attached at O-6 that may be further substituted at
O-2 by β-D-Galp units (Buckeridge, 2010). Besides that, a variety of
sugars, such as fucose, arabinose and galacturonic acid can be found
linked to the α-D-Xylp residues and the β-D-Galp units may be Oacetylated at different positions (Hsieh & Harris, 2009; Kiefer, York,
Darvill, & Albersheim, 1989).
Pectic polymers are defined as a class of polymers rich in galacturonic acid (GalA) and are usually water soluble. Three major pectic
polymers
are
recognized:
homogalacturonans
(HG),
Rhamnogalacturonans I (RGI) and Rhamnogalacturonans II.
Homogalacturonan is a linear polymer consisting of (1→4)-linked α-DGalpA, whilst rhamnogalacturonan I consists of the repeating disaccharide [→4)-α-D-GalpA-(1→2)-α-L-Rhap-(1→] to which a variety of
different glycan chains (arabinan, galactan and/or arabinogalactans)
are attached to the Rha residues. Finally, rhamnogalacturonan II has a
backbone of HG rather than RG, with complex side chains attached to
the to the GalpA residues (Willats et al., 2006).

Corresponding author at: CP 19.046, CEP 81.531-980 Curitiba, PR, Brazil.
Corresponding author at: 745 Agriculture Mall Drive, West Lafayette, IN 47907, USA.
E-mail addresses: (L.M.C. Cordeiro), (B.R. Hamaker).

⁎⁎

/>Received 29 May 2018; Received in revised form 22 October 2018; Accepted 6 November 2018
Available online 10 November 2018

0144-8617/ © 2018 Elsevier Ltd. All rights reserved.


Carbohydrate Polymers 206 (2019) 389–395

T. Moro Cantu-Jungles et al.

containing 50 mM 4-methyl-valeric acid (used as an internal standard
for SCFA analysis, No. 277827- 5 G, Sigma-Aldrich Inc., St. Louis, Mo.,
USA), meta-phosphoric acid (5%) and copper sulfate (1.56 mg/mL);
mixed and centrifuged at 13,000 rpm for 10 min. An aliquot (0.2 μL)
was injected into a GC-FID 7890 A (Agilent Technologies, Inc., Santa
Clara, CA, USA) equipped with a fused silica capillary column
(NukolTM, Supelco No. 40369-03 A, Bellefonte, Pa., USA). The initial
oven temperature was held at 50 °C for 2 min, ramped to 70 °C at a rate
of 10 °C/min, to 85 °C at a rate of 3 °C/min, to 110 °C at a rate of 5 °C/
min, to 290 °C at a rate of 30 °C/min, and finally held at 290 °C for
8 min. Helium was used as a carrier gas at a constant flow rate of 1 mL/
min through the column. Quantification was performed based on relative peak area of each short chain fatty acid in a fatty acid external
standard (Volatile Free Acid Mix, 10 mM, 46975-U, Supelco), adjusting
the quantity of each compound based on that of the internal standard.

Although ubiquitous in plants, there are no studies in the literature
comparing the gut fermentability of pectins vs xyloglucans, and in
forms commonly consumed (soluble pectins) vs uncommon (soluble
xyloglucans). Thus, we aimed to investigate the fermentation profile by
the human gut microbiota of previously isolated and characterized
pectins and a xyloglucan and investigate how gut commensal bacteria
differently respond to each polymer type.
2. Materials and methods

2.1. Dietary fibers
Two pectic polymers previously obtained and characterized by
Cantu-Jungles et al. (2014) were utilized: a HG together with a RGI
highly branched by type I arabinogalactans (HG + AGI) and the isolated RGI highly branched by type I arabinogalactans (AGI). Fraction
HG + AGI had a monosaccharide composition of Ara: Gal: Glc:GalA in a
23:21:2:54 ratio. The relative richness of HG versus AGI in HG + AGI
calculated based on the ratio of uronic acid/(Ara + Gal) was of 1.3 and
79% of the uronic acids in the homogalacturonan were methyl-esterified. Fraction AGI was composed by Ara: Gal: Rha: GalA in a ratio of
47.8:31.5:10.7:10 (Cantu-Jungles et al., 2014).
The xyloglucan was obtained from tucumã pulp as previously described (Cantu-Jungles, Iacomini, Cipriani, & Cordeiro, 2017). It was
mainly composed by Fuc:Xyl:Gal:Glc in a ratio of 6.2:19.4: 12: 62.4
(Cantu-Jungles, Iacomini et al., 2017).
Regarding the monosaccharide composition reported above for all
fractions, the uronic acid content is reported in weight %, while neutral
sugars are reported in molar amount.
Fructooligosaccharides (FOS - No. F8052, Sigma-Aldrich Inc.,
St.Louis, Mo., USA) was used as a positive control.

2.3. Bacterial shifts during the in vitro fecal fermentation
Metagenomic DNA was extracted from 1 mL of fecal ferments using
FastDNA SPIN Kit for Feces (MP Biomedicals, Laboratories, Carlsbad,
CA) according to the manufacturer's instructions. DNA integrity was
determined using a 1% agarose gel with ethidium bromide, and DNA
concentrations were verified using the Nanodrop ND-1000 (Nanodrop
Technology). DNA samples were stored at −20 C until sequencing.
Sequencing was performed at the DNA Services Facility at the
University of Illinois, Chicago. DNA was PCR amplified with primers
515 F and 806R (Caporaso et al., 2012), targeting the V4 variable region of bacterial and archaeal small subunit (SSU) ribosomal RNA
(rRNA) gene. Metagenomic sequencing was performed on an Illumina
MiSeq system. The identity of bacterial populations was analyzed using

QIIME software and R statistical software was used to carry out principal components analysis (PCA) (Caporaso et al., 2010).

2.2. In vitro fecal fermentation
3. Results
Batch fecal fermentation was performed as the methodology of
Lebet et al. (Lebet, Arrigoni, & Amadò, 1998) with some minor modifications (Rose, Patterson, & Hamaker, 2010). The total carbohydrate
in the samples was determined by the phenol-sulfuric acid method
(Dubois, Gilles, Hamilton, Rebers, & Smith, 1956). Each substrate
(50 mg equivalent carbohydrate) was weighed in 3 test tubes for triplicate analysis. Hydration of samples was performed by adding 4 mL of
carbonate-phosphate buffer pH 6.8 ± 0.1 to each tube. Fructooligosaccharides and tubes without any added carbohydrate were used as
positive and negative controls, respectively.
Fecal samples were obtained from 3 healthy volunteers who were on
their Western routine diet and had not taken antibiotics within the
previous 6 months. Fecal samples were collected in plastic bags that
were sealed after removing the air, transported on ice, and immediately
placed inside an anaerobic chamber (10% H2, 5% CO2, and 85% N2;
BactronEZ, SHEL LAB, Cornelius, OR) where all further procedures
were performed within 2 h after collection. Fecal samples were pooled
together, and the fecal slurry prepared by homogenization with carbonate-phosphate buffer pH 6.8 ± 0.1 in a ratio of 1:3 (w/v) and
further strained through 4 layers of cheesecloth. The hydrated carbohydrate sample and the controls were then inoculated with the filtrate
(1 mL). Tubes were sealed and incubated at 37 °C in a shaking water
bath. Immediately after incubation and after 4, 8, 12, and 24 h of fermentation, assigned tubes were removed from the water bath, and total
gas volume was measured by plunger displacement of the syringe.
Finaly, pH was measured and a copper sulfate solution (400 μL of
2.75 mg/mL) was then used as an antimicrobial agent, stopping microbial activity.

Previously obtained and characterized pectic polymers (AGI and
HG + AGI) and a xyloglucan (XYG) were submitted to an in vitro fecal
fermentation model. Profiles of generated metabolites (gas, pH and
SCFA) were analyzed at 4, 8 and 12 h of fermentation and compared to

the negative (Blank) and positive (FOS) controls. Fig. 1 shows the
amount of gas produced and pH alterations during in vitro fermentation.
Gas production was higher for pectic polymers than for the tested xyloglucan at 12 h fermentation. Drops in pH were also more intense
during fermentation of AGI and HG + AGI than the XYG. Moreover,
intense pH drops occurred up to 4 h in pectic ferments and FOS, while
pH drops in XYG lasted longer, up to the 8 h fermentation point (Fig. 1).
These are indicative that XYG presented a slower fermentation rate
compared to the tested pectins and FOS.
Regarding total SCFA, a similar production (61.6 ± 3.8 mM/50 mg
carbohydrate) was observed for pectic polymers and xyloglucan at 12 h
fermentation (Fig. 2). However, distinct SCFA were produced in higher
amounts depending on the polymer fermented. Fermentation of HG +
AGI led to the highest acetate production at 12 h, around 22% greater
than in XYG ferments. On the other hand, propionate was more effectively produced during XYG and AGI fermentation, resulting in a final
concentration ∼ 31% higher than that observed in HG + AGI ferments.
Both xyloglucan and pectic polymers led to low butyrate production
compared to the positive control (FOS) (Fig. 2).
To further explore bacterial preferences for each tested dietary fiber,
we performed DNA sequencing in the initial inoculum and all 12-hour
ferments (Figs. 3 and 4). In the initial inoculum and the blank, Firmicutes was the predominant phylum, making up 60 ± 1.3 and
56 ± 1.9% of the reads, respectively (Fig. 3). The abundance of Firmicutes was significantly reduced in all dietary fiber ferments tested
(Fig. 3). Conversely, the abundance of Bacteroidetes was significantly
higher in pectic and xyloglucan ferments, which presented a similar

2.2.1. Short chain fatty acid quantification
To quantify short chain fatty acid (SCFA) contents, aliquots (400 μL)
from fermented samples were combined with 100 μL of a mixture
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Fig. 1. Gas production (mL/50 mg carbohydrate) and pH variations during in vitro fecal fermentation of AGI, HG + AGI and XYG compared to FOS and the Blank.

Although total Bacteroidetes population was similar in all pectic and
xyloglucan ferments (Fig. 3), differences in community composition
emerged at lower phylogenetic levels (Fig. 4). Bacteroides plebeius was
higher in AGI (13.7 ± 2.4%) than HG + AGI (8.8 ± 2.4%) or XYG
(6.5 ± 0.8%) ferments (Fig. 4). On the other hand, Bacteroides

Bacteroidetes population of 47 ± 3.6, 41 ± 4.7 and 44 ± 1.4% for
AGI, HG + AGI and XYG, respectively, than in the initial inoculum and
the blank. Finally, in FOS ferments, a greater abundance of Actinobacteria was observed compared to the initial inoculum and the blank
(9 ± 0.3%) (Fig. 3).

Fig. 2. Total SCFA, acetate, propionate and butyrate production (mM/50 mg carbohydrate) during in vitro fecal fermentation of AGI, HG + AGI and XYG compared to
FOS and the Blank.
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Fig. 3. Composition of different bacterial phyla of samples at 12 h fermentation as determined by 16S rRNA gene sequencing. Only taxa greater than 0.8% total read
composition were included in this graph.

In the Firmicutes phyla relevant differences were found in some

bacteria from Clostridiales order that were significant higher in XYG
ferments (3.2 ± 0.1%) than in pectic ferments (2.4 ± 0.1% in both
AGI and HG + AGI) (Fig. 4). Within Clostridiales, some bacteria from
Lachnospiraceae family were also higher in XYG (15.3 ± 0.4%) than
pectic ferments (11.8 ± 1.1% in AGI and 12.2 ± 1.5% in HG + AGI).
Conversely, Blautia, a genus that also belongs to the Lachnospiraceae

uniformis population was highly stimulated in detriment of other Bacteroides species in the presence of XYG, but not pectic ferments. Indeed,
B. uniformis was the predominant specie in XYG ferments constituting
20 ± 0.7% of the total reads, a value 10 times higher than its population in the initial inoculum. Parabacteroides distasonis was also significantly higher in XYG (3.7 ± 0.3%) than pectic ferments
(1.2 ± 0,1% in AGI and 1.0 ± 0.2% in HG + AGI) (Fig. 4).

Fig. 4. Microbial composition of samples at 12 h fermentation as determined by 16S rRNA gene sequencing. Only taxa greater than 1% total read composition were
included in this graph.
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Fig. 5. Heat map analysis from 16S rRNA gene sequencing of the initial inoculum and fiber ferments at 12 h.

explained 83.1% of the variations between the different bacterial
communities. Both the analyses indicated that great bacterial community shifts occurred during xyloglucan fermentation but were not so
intense in pectic polymers fermentation. The latter, as well as the positive control (FOS) ferments, preserved a microbial composition more
similar to that of the original inoculum and the blank than xyloglucan
ferments (Figs. 5 and 6).

family, was higher in pectic ferments (6.5 ± 0.6% in AGI and

5.4 ± 0.3% in HG + AGI) than XYG (4.1 ± 0.5%) (Fig. 4). While a
similar bacterial composition in most Firmicutes could be observed
within pectic ferments, Lachnospira genera was significantly higher in
HG + AGI (4.1 ± 1.4%) than AGI (0.6 ± 0.0%) (Fig. 4).
Heat map and PCA analysis were conducted to investigate general
relationships among the bacterial communities from the different
dietary fiber ferments (Figs. 5 and 6). Hierarchical differences in heat
map analysis showed that while the initial inoculum, blank, FOS,
HG + AGI and AGI ferments samples formed one major cluster, the
xyloglucan ferments were present in a secondary, distinct cluster
(Fig. 5). In PCA analysis (Fig. 6), initial inoculum, blank, FOS, HG +
AGI and AGI ferments and XYG were distinctly separated by the first
principal component axis (PC1), rather than the PC2, which accounted
for 64.9% of the total variations. PC2 accounted for only 18.2% of the
variance in the bacterial communities. Overall, the PC1 and PC2 axes

4. Discussion and conclusions
Structural features of a polymer such as the molecular weight, the
type of monosaccharides released after digestion, the degree of polymerization and substitution, and other physical features that allows the
access of bacterial enzymes to the saccharides units may influence the
fermentation profile of a fiber in the colon, including its rate of digestion (Hamaker & Tuncil, 2014; Holscher, 2017). Thus, more complex
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Wolever, 2004) and its contribution to increase propionate in AGI ferments cannot be discarded.
The distinct metabolites profiles generated by each dietary fiber

tested, the microbial communities also varied accordingly, especially at
low phylogenetic levels. Indeed, shifts in gut microbiota composition
promoted by dietary fiber fermentation were shown to occur in very
specific species or strains (Chung et al., 2016). In our study, B. plebeius,
P. distasonis and bacterial groups such as Blautia, Lachnospira, Clostridiales and Lachnospiraceae, presented a distinct abundance on each
dietary fiber ferment. Moreover, a substantial increase in B. uniformis
population to the detriment of other Bacteroides species was evident in
XYG ferments. B. uniformis is among the most abundant organisms in
the microbiota of individuals in Western countries, and may be beneficial as it has been shown to exhibit the potential to ameliorate metabolic and immune dysfunctions associated with obesity (FernándezMurga & Sanz, 2016; Gauffin Cano, Santacruz, Moya, & Sanz, 2012). A
work of Larsbrink et al. (2014) showed that xyloglucans can be used as
energy source by most of B. uniformis strains in pure cultures. Our results reveal that even in the competitive environment of the fecal
complex microbiota, these xyloglucans were efficiently fermented by B.
uniformis.
Within pectic polymers (AGI and HG + AGI) only minor differences
in few bacterial groups were observed depending on the chemical
structure of the pectic polymer fermented. While Bacteroides plebeius
was higher in AGI, Lachnospira genera was higher in HG + AGI. This
data corroborates previous studies, which showed that some bacteria
such as Faecalibacterium prausnitzii and Bacteroides spp. respond differently depending on the pectin source and chemical structure
(Hamaker & Tuncil, 2014; Lopez-Siles et al., 2012).
PCA and heat map analysis indicate that pectic polymers lead to
microbial composition that resembles more of the initial inoculum than
the XYG ferments. Pectic polymers are present in its natural soluble
form in a variety of foods such as fruits and vegetables usually present
in a healthy diet (Wikiera, Irla, & Mika, 2014). The common presence of
soluble pectin in the diet could explain the higher similarity in composition of the microbiota of pectic ferments to the initial inoculum.
Xyloglucans, however, although ubiquitous in foods, are usually present embed in the plant’s cellulosic cell wall matrix (Hsieh & Harris,
2009; Scheller & Ulvskov, 2010) that may be difficult the access to
bacterial fermentation. In our study, the xyloglucan was isolated with
alkali and was applied in the fermentation experiment in its water-soluble form (Cantu-Jungles, Iacomini et al., 2017), and was accessible to

bacteria. This is an uncommon form of xyloglucans, and it is plausible
that their fermentation would drive to a very distinct fermentation
profile different from initial inoculum.
Overall, the results presented indicate that the pectic polymers and
xyloglucan, during their in vitro fermentation, drive different characteristic metabolites profiles and bacterial communities’ profiles. Such
an understanding of how isolated dietary fibers are fermented and affect microbial composition will be key to design new food ingredients
capable of modulating the intestinal microbiota in a targeted way.

Fig. 6. Principal component analysis (PCA) from 16S rRNA gene sequencing of
the initial inoculum and fiber ferments at 12 h.

polymers are expected to be slower fermented than simple structures.
The delayed fermentation profile of tested dietary fibers in this study
(xyloglucan > > pectins > FOS) supports this hypothesis. A slower
fermentation rate indicates that SCFA are prone to be delivered in all
portions of the large intestine, including the distal colon. The absence of
fermentable carbohydrates in more distal parts of the large intestine,
may lead to bacterial fermentation of proteins and consequent generation of detrimental metabolites such as hydrogen sulfide, phenols,
and ammonia (Blachier, Mariotti, Huneau, & Tome, 2007; MacFarlane,
Gibson, & Cummings, 1992). Thus, one could speculate that xyloglucan
and tested pectins should be able to exert a positive effect with fermentation occurring more distally in the colon in comparison to FOS.
Regarding short chain fatty acid production, each dietary fiber
produced a unique metabolite profile during in vitro fecal fermentation.
While FOS was highly butyrogenic, HG + AGI was more acetogenic,
and the XYG and AGI more propiogenic. While fermentation results of
xyloglucans is lacking in the literature, it was previously demonstrated
that pectic polymers from different sources are extensively fermented
by the colonic gut microbiota in vitro (Cantu-Jungles, Cipriani,
Iacomini, Hamaker, & Cordeiro, 2017; Gulfi, Arrigoni, & Amadò, 2005;
Jonathan et al., 2012; Licht et al., 2010; Min et al., 2015; Titgemeyer,

Bourquin, Fahey, & Garleb, 1991). However, these studies were conducted with the homogalacturonan (HG) alone (Cantu-Jungles, Cipriani
et al., 2017; Gulfi et al., 2005; Jonathan et al., 2012) or non-characterized pectic structures (Licht et al., 2010; Min et al., 2015;
Titgemeyer et al., 1991). In many plant types, however, type I rhamnogalacturonans branched by arabinans, galactans and/or type I arabinogalactan (AGI) can be found together with portions of HG (Willats
et al., 2006). Here we have shown that a HG together with a rhamnogalacturonan I highly branched by type I arabinogalactan (HG + AGI),
was specifically acetogenic with a low butyrate production similar to
what was found in other reports of HG alone (Cantu-Jungles, Cipriani
et al., 2017; Gulfi et al., 2005; Jonathan et al., 2012). However, when
AGI was isolated from HG, with butyrate production kept low, acetate
production was reduced, and propionate production was raised.
Aguirre, Bussolo de Souza, and Venema, (2016) also have shown that
fermentation of type II arabinogalactan (AGII) produced high propionate rates in the microbiota from obese subjects, similar to what was
found in our study for rhamnogalacturonan I branched by type I arabinogalactan (AGI). Indeed, in a study using mono and disaccharides,
Mortensen, Holtug, and Rasmussen, (1988) showed that propionate
production occurs during in vitro fecal incubation with a variety of
sugars, including arabinose, which is present in both type I and type II
arabinogalactans. Moreover, ingestion of L-rhamnose, a sugar present
in the main chain of RGI to which AGI is attached, was shown to increase propionate in the serum of healthy volunteers (Vogt, Pencharz, &

Acknowledgments
This research was supported by Projeto Universal (Process 477971/
2012-1) provided by CNPq foundation (Brazil), by PRONEXCarboidratos and CAPES. The author Thaisa Moro Cantu-Jungles received from CNPq Foundation a fellowship for a year of study at
Whistler Center for Carbohydrate Research at Purdue University (process 208166/2014-9) and for posdoctoctoral studies (Process 150235/
2017-8).
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